The discovery of the shrimp nephrocomplex (revision of the ...

Laboratory of Virology

Faculty of Veterinary Medicine

Ghent University

The discovery of the shrimp nephrocomplex (revision

of the antennal gland) and its role during WSSV-entry

and the moulting process

Gaëtan M.A. De Gryse

Dissertation submitted in fulfillment of the requirements for the degree of

Doctor in Veterinary Sciences (PhD)

2021

Promoters:

Prof. dr. Hans Nauwynck

Prof. dr. Peter Bossier

'Tis a lesson you should heed--

Try again;

If at first you don't succeed,

Try again.

Then your courage should appear;

For if you will persevere,

You will conquer, never fear,

Try again.

Once or twice though you should fail,

If you would at last prevail,

Try again.

If we strive,

'tis no disgrace

Though we did not win the race--

What should you do in that case?

Try again.

If you find your task is hard.

Try again;

Time will bring you your reward,

Try again;

All that other folk can do,

Why with patience should not you?

Only keep this rule in view,

Try again

William E. Hickson (and many more)

Print: University Press

Cover artist: Grydega

Promotors

Prof. dr. H. Nauwynck

Faculteit Diergeneeskunde, UGent, Belgium

Prof. dr. P. Bossier

Faculteit Bioingenieurswetenschappen, UGent, Belgium

Members of the examination committee

Prof. dr. W. Van Den Broeck

Voorzitter van de examencommissie Faculteit Diergeneeskunde, UGent, Belgium

Em. Prof. dr. P. Sorgeloos

Faculteit Bio-ingenieurswetenschappen, UGent, Belgium

Prof. dr. G. Smagghe

Faculteit Bio-ingenieurswetenschappen, UGent, Belgium

Em. Prof. dr. J. Vlak

Laboratory of Virology, Wageningen University, the Netherlands

Dr. V. Alday-Sanz

National Aquaculture Group of Saudi Arabia (Naqua), Saoudi-Arabia

Dr. J. Dantas Lima

Imaqua, Belgium

Table of Contents

7 General Introduction ............................................................................................. 15

1.1. Situating and importance ........................................................................... 17

1.2. The White spot syndrome virus ................................................................. 19

1.2.1. History of a global pandemic .............................................................. 19

1.2.3. Taxonomy .......................................................................................... 22

1.2.4. Viral structure ..................................................................................... 23

1.2.5. Host range and vectors ...................................................................... 28

1.2.6. Pathogenesis ..................................................................................... 29

1.3. General morphology of the penaeid shrimp ............................................... 37

1.4. The antennal gland .................................................................................... 39

1.4.1. Introduction ........................................................................................ 39

1.4.2. Functions of the antennal gland ......................................................... 40

1.4.3. Coelomosac ....................................................................................... 43

1.4.4. Labyrinth ............................................................................................ 45

1.4.5. Terminal duct ..................................................................................... 45

1.4.6. Overall morphology ............................................................................ 45

1.5. References ................................................................................................ 47

Chapter 2 Aims and Outline of the thesis ................................................................ 67

Chapter 3 Anatomical and morphological exploration of the antennal gland ........... 73

3.1. Abstract ..................................................................................................... 75

3.2. Introduction ................................................................................................ 76

3.3. Materials & methods .................................................................................. 78

3.3.1. Ex vivo 3D-reconstruction of HE-slices with AMIRA ........................... 78

3.4. Results ...................................................................................................... 80

3.4.1. General overview of the antennal gland morphology .......................... 80

3.4.2. Compact glandular compartment ....................................................... 82

3.4.3. The rostral compartment .................................................................... 84

3.4.4. The ventral bladder ............................................................................ 86

3.4.5. The median compartment................................................................... 87

3.4.6. The lateral compartment with the caudal extensions .......................... 88

3.5. Discussion ................................................................................................. 90

3.5.1. Anatomy of the antennal gland based on HE-staining ........................ 90

3.5.2. The flow of filtrate to and through the antennal gland ......................... 92

3.5.3. Muscles associated with the antennal gland and their functions......... 94

3.5.4. Possible mechanical role of the antennal gland in moulting ............... 95

3.6. Conclusion ................................................................................................. 96

3.7. References ................................................................................................ 97

Chapter 4 µMRI imaging of Penaeus vannamei: the involvement of the shrimp

nephrocomplex during the moulting process ......................................................... 101

4.1. Abstract ................................................................................................... 103

4.2. Introduction .............................................................................................. 104

4.3. Material and methods .............................................................................. 105

4.3.1. In vivo 3d-reconstruction of the antennal gland using µMRI ............. 105

4.4. Results .................................................................................................... 106

4.4.1. Immobilisation of shrimp by lowering the ambient temperature ........ 106

4.4.2. Differentiation between nephrocomplex and surrounding tissues ..... 106

4.4.3. In vivo confirmation of the nephrocomplex morphology .................... 107

4.4.4. Volume shifts during different moulting stages ................................. 110

4.5. Discussion ............................................................................................... 111

4.5.1. µMRI ................................................................................................ 111

4.5.2. Moulting............................................................................................ 112

4.6. Conclusion ............................................................................................... 113

4.7. References .............................................................................................. 114

Chapter 5 Nephropore structure and dynamics during mictio ................................ 117

5.1. Abstract ................................................................................................... 119

5.2. Introduction .............................................................................................. 120

5.3. Material and methods .............................................................................. 120

5.3.1. Scanning electron microscopy of the Penaeus vannamei nephropore

................................................................................................................... 120

5.3.2. Histology .......................................................................................... 121

5.3.3. Determination of nephropore opening pressure in an ex vivo model 122

5.4. Results .................................................................................................... 125


................................................................................................................... 125

5.4.2. Histological examination of the nephropore ...................................... 125

5.4.3. Determination of nephropore opening pressure in an ex vivo model 126

5.4.4. Determination of fluid and particle influx in an ex vivo model after

simulated urination. .................................................................................... 126

5.5. Discussion ............................................................................................... 127

5.6. Conclusions ............................................................................................. 130

5.7. References .............................................................................................. 131

Chapter 6 Artificial and natural infection of the nephrocomplex ............................. 133

6.1. Abstract ................................................................................................... 135

6.2. Introduction .............................................................................................. 136

6.3. Material and Methods .............................................................................. 137

6.3.1. artificial infection ............................................................................... 137

6.3.2. Natural infection using salinity shock immersion .............................. 142

6.4. Results .................................................................................................... 142

6.4.1. Validation of intrabladder inoculation ................................................ 142

6.4.2. Artificial infection of the nephrocomplex ........................................... 143

6.4.3. Natural infection of the nephrocomplex ............................................ 149

6.5. Discussion ............................................................................................... 149

6.6. Conclusions ............................................................................................. 150

6.7. References .............................................................................................. 151

Chapter 7 General Discussion .............................................................................. 155

7.1. The shrimp’s nephrocomplex (antennal gland) is much more complex than

previously assumed. ....................................................................................... 157

7.2. The shrimps nephrocomplex is involved in the moulting process ............ 159

7.3. The shrimp’s nephrocomplex is a major entry portal of pathogen entry ... 160

7.4. The secondary defences of the shrimps nephrocomplex ......................... 165

7.5. Implications for Science and the industry................................................. 167

7.6. Conclusions ............................................................................................. 168

7.7. References .............................................................................................. 169

Summary............................................................................................................... 175

Samenvatting ........................................................................................................ 183

Curriculum Vitae and publications ......................................................................... 191

Acknowledgements-Dankwoord ............................................................................ 199

List of relevant abbreviations 3D Three dimensional

A Early post-moulting stage

B Late post-moulting stage

C Intermoult

CA Coxipodite adductor

CE Caudal extensions

cfu Colony forming units

CGG Compact glandular compartment

D1 Early pre-moulting stage

D2 Late pre-moulting stage

DL Dorsal lobe of the median compartment

DNA Deoxyribonucleotide

FAO Food and Agricultural Organisation

HE Haematoxylin and eosin staining

hpi hours post infection

IIF Indirect immunofluorescence

IM Intramuscular

LC Lateral compartment

LD50 Lethal dose 50%

LSA Large scaphocerite adductor

MBW Mean body weight

MC Median compartment

µCT micro computational tomography

µMRI Micro magnetic resonance imaging

NC Nucleocapsid

NP Nephropore

PEC pre-esophageal connection

PO Per oral

PoEC Post-esophageal connection

PVC Polyvinyl chloride

qPCR quantitative Polymerase chain reaction

RC Rostral compartment

SEM Scanning electron microscopy

SID50 Shrimp infectious dose 50%

SPF Specific pathogen free

SSA Small scaphocerite adductor

VB Ventral bladder

VL Ventral lobe of the median compartment

VP Viral protein

WSSV white spot syndrome virus

Chapter 1

General Introduction

G.M.A. De Gryse

Department of Virology, Parasitology and Immunology Faculty of Veterinary Medicine

Ghent University, Merelbeke, Belgium

Chapter 1 General Introduction

17

1.1. SITUATING AND IMPORTANCE

The exponentially rising global population is a dire problem facing humanity today. The global

human population is predicted to reach over 9 billion people in 2050 (Golden et al., 2016).

Although the European population is likely to decrease, the number of inhabitants in

developing countries is expected to grow enormously (UN, 2013). Overpopulation raises

paramount challenges such as the prevention of worldwide food insecurity. In 2017, already

821 million people are estimated to be chronically malnourished (FAO et al., 2018). To

maintain this growing world population, the food production must expand accordingly.

Agriculture needs to fill that gap. However, the FAO reports that there is less than 12% usable

land left for agriculture (FAO, 2008). Doubling the workable ground surface is a possibility,

but requires a mass destruction of forest, loss of biodiversity, and carbon sequestration

capacity. Furthermore, there is the question of freshwater supply. In 2012, already 70% of the

available 3,800 km3 of freshwater is being used by agriculture and 20% by industry (FAO,

2012). The growing population will need double the amount of freshwater it consumes now

and the necessary agricultural increase combined with the climate change will further stress

freshwater supplies. Marine aquaculture in contrast, has the potential of providing the

required protein without competing for freshwater or workable land. Further developing the

marine aquaculture will therefore be part of a sustainable strategy to meet the growing need

of food production to secure food security for the rising global population.

Today, aquaculture is the fastest growing animal production industry in the world (Waite et

al., 2014). In 2018, total aquaculture production reached 114.5 million tons and in the period

2001-2018, the industry achieved an annual growth of 5.3%. Global aquaculture production

of shrimp reached 4.9 million tons, with a total value of €45 billion, while production in 2003

was still at 2 million tons (€7.52 billion). As a high-value species, shrimp production is

projected to grow even further in the future. As a comparison: the total production of salmon

clocked off at 2.3 million tons (FAO, 2012, 2020).

Experts agree that aquacultured seafood can and will play a decisive role in the quest for

global food security (Béné et al., 2015). However, the industry should than expand even more

than it already does today. This is hindered by production losses due to (infectious) diseases


18

(Stentiford et al., 2012; FAO, 2020). Industry wide, over 6 billion dollars are lost annually due

to these infectious diseases (World Bank, 2014). Already, many aquaculture branches took

huge leaps towards profound progress in identifying, diagnosing, treating, and managing

diseases (Groner et al., 2016). The shrimp aquaculture industry in contrast has not yet

managed to adequately tackle infectious diseases, often leading to socio-economic disasters

on local as well as global scale. Up to 80% of losses are caused by pathogens of shrimp (Flegel

et al. 2008; Stentiford et al. 2012). To illustrate the importance: the World Organisation of

Animal Health (OIE) lists 116 notifiable diseases of which eight are shrimp related, namely

white spot syndrome virus (WSSV), yellow head virus (genotype 1, YHV1), Taura syndrome

virus (TSV), nodavirus (white tail disease), infectious myonecrosis virus (IMNV), infectious

hypodermal and haematopoietic necrosis virus (IHHNV), Hepatobacter penaei (necrotising

hepantopancreatitis) and acute hepatopancreatic necrosis disease (AHPND). Six of these are

viruses and only two bacteria (OIE 2020). Of all these viruses, the WSSV is undoubtedly the

most notorious and destructive. This virus is the main cause of losses in global shrimp

production as all cultured penaeids, plus most other Crustacea are susceptible to WSSV (see

1.2.4) (Walker and Mohan, 2009; Pradeep et al., 2012).

The main route of WSSV transmission is still a blind spot on the WSSV mitigation map. The

natural main entry portal for WSSV through which WSSV can enter the shrimp and reach the

primary replication site is still unknown. Consequently, current knowledge on pathology,

pathogenesis and possible virus inhibition is almost exclusively based on studies where the

animal was infected by injection, which is not a natural route of infection and possibly

confounds conclusions. Identifying the WSSV entry portal is therefore necessary for the

advancement of research and more specific, prevention and treatment measurements.

Investigating the possible weaknesses in the barriers of shrimp, from which the virus could

benefit, is therefore of paramount importance.


19

1.2. THE WHITE SPOT SYNDROME VIRUS

1.2.1. History of a global pandemic

In 1992 a new disease emerged in northern Taiwan. Reports mention the apparition of white

spots on the cephalothorax followed by mass mortality in Penaeus japonicus. The next year

the same symptoms were seen in Taiwanese Penaeus monodon and Penaeus penicilatus

farms, causing great losses (Chou et al., 1995). Next, the unknown disease crossed the East

China Sea to assert its effects on Chinese shrimp farms with heavy losses as consequence

(Zhan et al., 1998). In March 1993, this disease - meanwhile named white spot syndrome after

the then pathognomic white spots - reached Japan via imported Chinese P. japonicus juveniles

(Nakano et al., 1994). It was from this outbreak that the causative agent, a virus, was

identified and visualized for the first time (Inouye et al., 1994). In the following years, the

WSSV-pandemic spread over at least 42 countries, throughout all continents (Chou et al.,

1995; Calderón et al., 1999; APHIS-USDA, 2004; Eissa et al., 2009; Stentiford and Lightner,

2011). Figures 2, 3 and 4 show the distribution of WSSV over global waters.

Figure 2. Emergence of White spot syndrome throughout the world from 1992 to 1994. 1992: Taiwan (Chou et

al., 1995). 1993: China (Zhan et al., 1998), Japan (Nakano et al., 1994), Korea (Park et al., 1998), Vietnam (Khoa

et al., 2001). 1994: Thailand (Lo, 1996), India (Wongteerasupaya et al., 1995), Bangladesh (Mazid and Banu,

2002).

Thailand is thought to be the main hub for WSSV-migration across Southeast Asia. Based on

genetic differences in the open reading frame (ORF) 14/15, the TH-96-II strain is probably the


20

ancestral strain of the WSSV-TH strain (both from Thailand). WSSV-TH eventually mutated

into three other strains: (i) WSSV-TW (Taiwan) that evolved into WSSV-CN (China), (ii) WSSV-

VN (Vietnam) and (iii) two Indian strains IN-05-I and IN-07-I (Pradeep et al., 2008).

The enormous potential of the virus to rapidly disseminate to several other nations is

demonstrated by the history of the continental spread in America. In mid-January 1999, WSSV

first arrived in Central America and was almost immediately detected in Nicaragua, Honduras,

and Guatemala (Walker and Mohan, 2009). In April 1999, only a few months later, the virus

reached Panama. Ecuador reported WSSV positive cases in May, followed by Peru in August

and Colombia by the end of 1999. Costa Rica and Mexico followed shortly in 2000 even though

Mexico had implemented preventive measures against the WSSV-threat (de Graindorge and

Griffith, 2000; Hill, 2002; Mialhe et al., 2013). The description of this spread was reconstructed

based on early warning reports. It is possible that WSSV was already present in those

countries at the time of first detection. The outbreaks of WSSV followed a cold storm weather

moving along the coast. In 1999, 25% of Penaeus vannamei post larvae (wild and hatchery)

were WSSV-positive on histology samples from as far as 1996. At the time, Panama was the

main distributor of post larvae to the rest of Latin-America (Victoria Alday-Sanz, personal

communication).


21

Figure 3. Spread of the White spot syndrome virus around the world from 1995 to 2002. The Brown coloration

is displayed in more detail in Figure 2. 1995: Greece (Stentiford and Lightner, 2011), Sri Lanka (Siriwardena,

2001), USA (APHIS-USDA, 2004), Malaysia, Indonesia (Kasornchandra et al., 1998). 1996: Cambodia (Zwart et al.,

2010). 1997: Italy, Turkey (Stentiford and Lightner, 2011). 1999: Honduras, Nicaragua, Guatemala (Hossain et

al., 2001), Panama (Rosenberry, 1999), Ecuador (Calderón et al., 1999), Philippines (Magbanua et al., 2000),

Colombia (OIE, 1999), Peru (Mialhe et al., 2013), Pakistan* (Bondad-Reantaso et al., 2001), Taipei** (OIE, 1999).

2000: Togo (OIE, 2000), Mexico (Bondad-Reantaso et al., 2001), Australia (East et al., 2004), Spain (Stentiford

and Lightner, 2011), Costa Rica (Vargas, 2001). 2002: France, Iran (OIE, 2002), Myanmar (FAO/NACA, 2003). (*)

Infection with WSSV was suspected but not confirmed. (**) Taipei was not indicated on the map because of the

small land area.

The swift global spread as shown in Figures 2, 3 and 4 of the WSSV virus was only possible

because of the several ways WSSV can enter a country. The intensive (post)juvenile shrimp

trade ensured a rapid distribution of the virus within Southeast Asia (Zhan et al., 1998; Flegel,

2006). The import of frozen crustaceans as food for other farmed crustaceans also played a

significant part in the introduction of WSSV in shrimp farms (East et al., 2004). Furthermore,

farmed shrimp may escape from aquaculture ponds and facilities and reach the open sea,

either via wastewater or by flooding during the rainy season. If these escaped shrimps are

infected, they can pass the WSSV virus to the wild crustacean population. Both the wild and

escaped crustaceans can be the cause of further dissemination to other farms and countries.

Additionally, the use of imported (frozen) shrimps as bait in sport fishing is also suggested to

lead to WSSV-introduction in naïve wild populations. Moreover, it has been described that

the larvae of shrimp are taken up by ships via the ballast water and doing so, travel greater

intercontinental distances (Lightner et al., 1997; Flegel, 2002; Fuller et al., 2014). Captive

breeding farms catch shrimp from the wild population to replenish their breeding stock or

use as new breeding material. This way, the virus can enter the farm and infect parallel

production lines (Flegel, 2006). Hsu et al. (1999) used a wild population of P. monodon as a

breeding stock for experimental studies and found that only 33 % was negative for WSSV.

Also, wastewater from food processing plants can carry the virus to wild populations and

other farms. For example, a human food processing plant that imported frozen shrimp from

Asia most probably caused the WSSV introduction in Texas at the end of 1995 (Lightner et al.,

1997). These transmission pathways demonstrate that effective prevention against WSSV

spread should not only be limited to the aquaculture specific measures alone.


22

Figure 4. Further spread of the White spot syndrome virus from 2003 to 2011. The Brown coloration is displayed

in more detail in Figures 2 and 3. 2003: Argentina (Martorelli et al., 2010), Hong Kong* (OIE, 2003). 2004: El

Salvador (OIE, 2004). 2005: Brazil (APHIS-USDA, 2005). 2009: Egypt (Eissa et al., 2009). 2010: Saudia Arabia,

Madagascar, Mozambique (Tang, 2013). 2011: Brunei Darussalam (OIE, 2011). (*) Hong Kong was not indicated

on the map because of the small land area.

1.2.3. Taxonomy

The classification of WSSV has long been the cause of discussion and confusion, starting from

the very first article on electron microscopic imaging of the virus by Inouye et al. (1994).

Inouye et al. (1994) and his collaborators temporarily coined the virus "Rod-shaped nuclear

virus of Penaeus japonicus". Over the years, several different names were given to the virus

as displayed in Table 1 (Escobedo-Bonilla et al., 2008). Finally, in 1996, a consensus was

reached, and the virus was renamed ‘White spot syndrome virus’ (Lightner, 1996a; Escobedo-

Bonilla et al., 2008). However, confusion remained for several years as many publications kept

using different names for the same virus.


23

Table 1. Overview of the different names appointed to the White spot syndrome virus.

Name Abbreviation Reference Rod-shaped nuclear virus of Penaeus japonicus RV-PJ Inouye et al., 1994 Third Penaeus monodon non-occluded baculovirus

PmNOB III

Wang et al., 1995

White spot syndrome baculovirus WSSV Wang et al., 1995 White spot baculovirus WSBV Chou et al., 1995 Systemic ectodermal and mesodermal baculovirus

SEMBV

Wongteerasupaya et al., 1995

Hypodermal and haematopoietic necrosis baculovirus

HHNBV

Jie et al., 1995

Penaeid rod-shaped DNA virus PRDV Venegas et al., 2000 White spot syndrome virus WSSV Lightner, 1996

Due to the virus’ morphology (see 1.2.3.), consensus first classified the WSSV within the family

of the Baculoviridae (see table 1). However, genomic analysis uncovered that WSSV is a virus

from a previously unknown genus and family, now named Whispovirus and Nimaviridae

respectively. Up till the publication of this thesis (2021), WSSV is the only representative of

this genus, but in 2019, five novel nimaviridae family-members were uncovered. It does not

belong to any order or subfamily (Van Hulten and Vlak, 2002; ICTV, 2012; Kawato et al., 2019).

1.2.4. Viral structure

When observed under the electron microscope with negative staining, the WSSV virion can

be observed as cylindrical to ellipsoidal construction with a filamentous appendix (270-310

nm x 30 nm), similar to the shape of a spermatozoid (Figure 5) (Wongteerasupaya et al., 1995;

Durand et al., 1996). The width of an intact virion, including the envelope, varies between 70

and 167 nm and is 250 to 420 nm long (Table 2). However, the visually most striking feature

of the virion is its long filamentous appendix. It served as an inspiration in opting the family

name ‘Nimaviridae’ ("nima" is Latin for "thread or rod"). The function or composition of this

appendix has not yet been unraveled (Dantas-Lima, 2013). During a study of the viral

replication cycle, it was hypothesised that the tail is a remnant of the budding process in the

nucleus of infected cells (Li et al., 2015). Furthermore, a plug-like structure is present at the

annexation place of this 'tail', which is in fact an extension of the envelope (Figure 5A) (Tsai

et al., 2006). Electron microscopical images (Tsai et al., 2004; Tsai et al., 2006) show that the


24

envelope is a 6-8 nm thick structure, consisting of a trilaminar lipid membrane. Each laminar

lipid layer of the membrane is in its turn bilaterally surrounded by an electron transparent

layer (Figure 5B) (Inouye et al., 1994; Durand et al., 1997; Kasornchandra et al., 1998). The

viral proteins (VP) 28 and 19 are expressed on the outside of the envelope (Tsai et al., 2004;

Tsai et al., 2006).


25

Table 2. Overview of the reported dimensions of White spot syndrome virus.

Intact virion*

(nm)

NC(+)

(nm)

NC(-)

(nm)

ΔNC

(nm)

Country of origin

L W L W L W L W

330 87 220 70 - - - - Taiwan1

330 120 275 85 - - - - China²

350-

375

150-

157 325 120 395 83 +70 - 37 China³

275 83 - - - - - - Japan4

275 83 216 54 - - - - Japan²

375 167 290 75 - - - - Korea5

276 121 201 89 302 73 +101 -16 Thailand6

250-

380

70-

150 - -

330-

350 58-67 - - Thailand7

300-

420

110-

140 - -

300-

420 70-95 - - Thailand8

275 120 236 85 - - - - Thailand²

266 112 - - 420 68 - - India 9

320 80-

100

182-

250 60-80 - - - - India 10

270 110 260 80 - - - - India ²

320 120 260 80 - - - - Indonesia²

275 120 230 85 - - - - Malaysia ²

NC(+): nucleocapsid surrounded by envelope, NC(-): nucleocapsid not surrounded by envelope, ΔNC: difference

in size of naked nucleocapsid (+ is larger and - is smaller). L: Length, W: Width. Lowest value, highest value.

References: 1Chou et al., 1995, 2Kasornchandra et al., 1998, 3Zhan et al., 1998, 4Inouye et al., 1994, 5Park et al.,

1998, 6Wongteerasupaya et al., 1995, 7Wang et al., 1995, 8Durand et al., 1996, 9Hameed et al., 1998, and 10Rajendran et al., 1999. (*) The filamentous appendix was not considered when determining the size of the

intact virion.


26

Underneath the trilaminar envelope, the nucleocapsid ellipsoid-shaped nucleocapsid with a

banded pattern can be observed. This pattern is obtained by an accumulation of ring-shaped

structures consisting of VP664 (Figure 5D & 5E) (Tsai et al., 2006). The wall of the nucleocapsid

is 6 nm thick (Hameed et al., 1998; Kasornchandra et al., 1998) and the core is highly electron-

dense, containing the DNA and DNA-binding protein VP15. As described, the nucleocapsid is

ellipsoid under normal circumstances. Per contra, during the replication cycle when the

envelope not yet enrobes the nucleocapsid, the nucleocapsid possesses a longer, narrower,

cylindrical aspect. The same conformation is observed when the envelope ruptures (Figure

5). The difference in non-enveloped (naked) and enveloped nucleocapsid length is seldomly

reported, but a considerable variation appears to be present. The naked nucleocapsid will

mostly expand without envelope so it becomes longer than the intact virion (Table 2 and

Figure 5E). These changes in shape and length indicate that the nucleocapsid is assembled

under tension inside the envelope, perhaps acting like a coil spring upon entering a host-cell

(Wongteerasupaya et al., 1995; Hameed et al., 1998; Zhan et al., 1998; Escobedo-Bonilla et

al., 2008). In addition, degrading nucleocapsids have a swollen and hatched appearance and

give an untensed impression compared to the above-described cylindrical shape of the

normal naked nucleocapsid. (Durand et al., 1996; Durand et al., 1997).


27

Figure 5. Structure of the White spot syndrome virus. 5A-Complete, intact virion (Computer enhanced). Ex:

filamentous extension, Env: envelope. P: Plug-like structure. 5B-Electron microscopical recording of an intact

virion. NC: nucleocapsid. 5C-Electron microscopical recording of an intact virion with an erratic implantation of

the filamentous extension (EEx). 5D-Electron microscopical recording of a naked nucleocapsid. The conformation

of the naked nucleocapsid is clearly distinguishable. The bandaged pattern is also very well visible here. White


28

arrow: ring-shaped protein structure. 5E-Electron microscopical recording displaying a visible clear length

difference between the naked nucleocapsid (white arrows) and intact virions (black arrows). Origin of pictures:

Tsai et al. (2006) (5A), Wongteerasupaya et al. (1995) (5B), Durand et al. (1997) (5C), Leu et al. (2005) (5D & 5E).

Finally, between the envelope and the nucleocapsid, there exists a third structure; the so-

called tegument. This tegument is a partially permeable layer through which the button-

domain of VP664 is penetrated. Viral protein 26 is the main protein of the tegument and is

also present on the outside of the tegument, where it interconnects with the protruding

VP664 domains. This conformation probably provides the required flexibility to get the

nucleocapsid inside the envelope during the replication cycle. Viral protein 26 dissolves in

high salt concentrations, causing the nucleocapsid to take on the above mentioned more

rigid, cylindrical form (Tsai et al., 2006).

1.2.5. Host range and vectors

In short, all Decapoda are suspected to be susceptible to WSSV infection including shrimp,

prawn, lobsters, crabs, and crayfish. Both cultivated and wild species are at risk (Stentiford et

al., 2009; Bateman et al., 2012; Pradeep et al., 2012). This includes major economical

significant European crustaceans such as Nephrops norvegicus, Homarus gammarus, and

Cancer pagurus, which are also described to be highly WSSV-susceptible (Bateman et al.,

2012; Pradeep et al., 2012; Stentiford et al., 2009). Furthermore, other Crustacea such as

important food sources in aquaculture like Artemia sp. and Copepoda and even other

invertebrates like polychaetes are also susceptible. Additionally, some insect larvae have

been found to contain WSSV (Desrina et al., 2013; Pradeep et al., 2012). For a more extensive

list of all sensitive species, see the reviews of Escobedo-Bonilla et al., (2008) and Pradeep et

al., (2012). Notwithstanding, a difference in sensitivity to disease and mortality exist within

the subphylum of the Crustacea. Especially species not belonging to the Penaeidae family are

less vulnerable and do not so easily succumb to the virus (Stentiford et al., 2009). Moreover,

the Macrobrachium rosenbergii has been thought to be completely refractive to WSSV

infection, unlike all other members of the Crustacea. Yet, under standardised conditions,

WSSV-replication does take place in M. rosenbergii and can potentially inflict mortality. Still,

Penaeus vannamei has been found to be 18.6 to 398 times more susceptible to WSSV-


29

infection than the economically less valuable M. rosenbergii (Corteel et al., 2012; Escobedo-

Bonilla et al., 2005). However, for most crustaceans, the capacity of WSSV to disease and

affect the population remains unknown. Here, observational research is direly needed on the

European species relevant for the fishery industry. Especially since the virus has already been

reported in Europe and was registered as non-exotic in EC Directive 2006/88. Case reports

show that WSSV can cause morbidity and mortality at European temperatures (Stentiford and

Lightner, 2011). Several susceptibility studies have demonstrated the virulence of WSSV in

native species, including Nephrops norvegicus (Norway lobster) and Crangon affinis (closely

related to the majorly important grey Northern prawn, Crangon crangon) (Gong et al., 2010;

Bateman et al., 2012). The value of the C. crangon fishery in the North Sea alone is estimated

at more than €100 million (Verhaegen et al., 2012). The potential to plunge the indigenous

European wildlife stock and the resulting impact on jobs, food supply, ecology and the

economy is therefore certainly something that must be considered (Hill, 2002). The need for

prospective studies on this potential threat is pressing. Especially in view of global warming

and shifting water temperature, WSSV-replication in local populations becomes more likely.

1.2.6. Pathogenesis

Symptomatology

When WSSV manages to invade a shrimp farm, the consequences are often imminent and

disastrous. If a naïve population is infected, two to 7 days after the first infected shrimp show

symptoms, a pond morbidity and mortality of both 100% is often observed. First, affected

shrimp become lethargic and their appendages discolour red to pink, they lose their appetite

and the cuticula begins to loosen. In the early days of the pandemic, white spots appeared on

the inside of the carapax due to calcium deposits (diameter from 0,5 mm to 1 cm). However,

these white spots, to which the virus owes its name, are no longer frequently observed today.

One to 2 days before mortality, shrimps gather at the edge of the pond before death sets in

(Chou et al., 1995; Chang et al., 1996; Lightner, 1996b; Lo, 1996; Kasornchandra et al., 1998;

Kou et al., 1998). Over the years, as WSSV became pandemic and enzootic in several shrimp

producing countries, the severity of WSSV-infection began to diminish (Stalinraj et al., 2009).

This is on its own proof that the shrimp population can adapt to viral selection pressure over


30

time and different generations. The theory of viral accommodation by Flegel could explain

how this feat is accomplished. By integration of WSSV-genetic material in the host genome

(so called endogenous viral elements, EVE), shrimp could establish resistance to these viruses

in a population by fortuitous production of antisense, immunospecific RNA (Flegel, 2007;

Utari et al., 2017). Especially when these elements are integrated in the reproductive cells.

However, it is not because some infected animals appear to show no overt symptoms, that it

can be concluded that WSSV-infection is without harm. Several reports describe less

productivity and reduced tolerance to stress and other infection in WSSV-endemic

populations (Debnath et al., 2014; Zhu et al., 2019), potentially making WSSV to shrimp

industry, what bovine viral diarrhoea and infectious bovine rhinotracheitis is to the cattle

industry (Gunn et al., 2004; Nettleton and Russell, 2017).

Latency and reactivation

Although clearly very pathogenic, WSSV can be latent in asymptomatic animals and was even

found in shrimp certified as specific pathogen free (SPF). The mechanism of latency and

reactivation of WSSV is not fully understood. In PCR-negative shrimp, DNA microarray

experiments exposed three ORFs that were strongly expressed (ORF 151, 366 and 427) while

structural genes such as VP28, VP24 and VP15 were relatively little expressed. In PCR-positive

asymptomatic shrimp, these conditions were reversed (Khadijah et al., 2003). ORF 89 and ORF

403 act similarly as later observed by Hossain et al. (2004) and He and Kwang (2008). It was

found that ORF 89 will suppress its own promoter as well as the WSSV-thymidine-thymidylate

kinase and protein kinase genes at the transcriptional level, effectively inhibiting the WSSV-

replication (Hossain et al., 2004). The ORF 403 codes for an E3 ubiquitin ligase and ORF 427

binds with a previously unknown serin/threonine protein phosphatase, thus propagating viral

latency (Lu and Kwang, 2004; He and Kwang, 2008). In conclusion, the presence of latent

WSSV coincides with very low transcription of structural genes VP28, VP24 and VP15, and

upregulation of genes encoded by WSSV ORF 89, 151, 366, 403, and 427. In reaction, the

shrimp will respond to this latency by expressing more phagocytosis activating protein

(Shekhar et al., 2012) and other proteins related to actin-myosin cytoskeleton process,

including myosin, actin, tubulin, clathrin, and tropomyosin, which are also involved in

phagocytosis (Li et al., 2016). Apoptosis inhibiting proteins were also found to be upregulated

during latent WSSV-infection i.e. ALG-2 interacting protein x, Heat shock protein 90, 14-3-3-


31

like protein, peroxiredoxin 5, peroxiredoxin 6 and adenine nucleotide translocase 2 (Li et al.,

2016). It was demonstrated that reactivation of latent WSSV is likely to occur during sudden

environmental changes such as an acute change in salinity (Liu et al., 2006) and/or

temperature (Jiravanichpaisal et al., 2004). In addition, it was suggested that spawning can

reactivate latent virus by a significant increase of the active signal-transduction-and-activator-

of-transcription-protein (STAT, due to phosphorylation) during and after spawning (Lin et al.,

2012). STAT activation is important for the transcription of the immediate-early gene ie-1 of

WSSV, in turn triggering the viral replication cascade (Liu et al., 2005; Liu et al., 2007). Latent

virus mainly resides in muscular tissue, where it has little to no effect on shrimp physiology.

Some researchers claim that shrimp production parameters will not be impaired by the

presence of a latent WSSV-infection, while others say that the animals become less resilient

to environmental stress and more susceptible to xenobiotic contamination (Zhu et al., 2019).

Finally, Debnath et al. (2014) did find a negative influence of WSSV-positive P. monodon

breeding animals on their mature eggs; of all mature eggs laid by infected brood stock, fewer

eggs hatched in comparison to eggs laid by WSSV-negative conspecifics.

Transmission and WSSV-entry

The WSSV-spread from country to country and from farm to farm was discussed earlier (see

above). How the virus spreads from shrimp to shrimp is not yet entirely clear. Nevertheless,

a profound understanding of the disease’s pathogenesis is one of the most crucial aspects in

reaching an adequate protection against pathogens, be it prophylactic, in management, or

curative. Without exceptions, all infectious diseases start with the very same step, i.e.,

pathogen entry and subsequent access to the primary replication site. Thus, knowing where

the infectious particles enter the animal body is crucial information. For example, if the entry

portal is known, then local defense mechanisms can be studied, upregulated, or induced (e.g.,

local immunity inducing vaccines, probiotics, etc.). Furthermore, this information provides

hints to possible risk factors, while management strategies can be undertaken to cut the chain

of transmission (e.g., animal isolation, feed supply, waste management, hygiene, vector

control, etc.). Finally, when the method of inoculation mimics the actual, natural way of

infection, further pathogenesis studies can be performed with higher accuracy. Based on the

general anatomy of shrimp (Young 1959), six potential pathogen entry portals can be

considered: (i) mouth (peroral) (ii) anus, (iii) the nephropore, (iv) integumental gland, (v) the


32

genital orifices and (vi) the entire surface of the body, when the exoskeleton is somehow

compromised (including eyes, gills, etc.).

Chang et al. (1996) demonstrated that the cells of the stomach, antennal gland,

hepatopancreas, gills and the cuticular epidermis were the first to be infected. Later, the

infection spreads to other organs such as the heart, intestines, hematopoietic tissue, and

lymphoid organ. A few hours later, virus was found in the eye, nervous system, and muscle

tissue. In short, it can be stated that all mesodermal and ectodermal cells of the shrimp are

susceptible to the virus (Lo et al., 1997; Escobedo-Bonilla et al., 2008). With so many cell-

types and species highly susceptible to WSSV and other pathogens, it can almost be called

miraculous that shrimp still manage to survive. The explanation can be found in the shrimp’s

ability to defend itself against pathogens, particularly in its primary defense system located

at the potential pathogen entry points with the main function of preventing pathogen entry.

The first, most evidently apparent defense system is the shrimp’s exoskeleton. This cuticular

layer is not only protective against physical threats but is also very effective in fending off

other noxes such as pathogens and toxins (Corteel et al., 2009; Xiong et al., 2017). The cuticula

is made of chitin fibers with protein deposits, reinforced by minerals such as calcium and

magnesium. This combination makes the structure strong and resilient, but also very dense.

Furthermore, significant parts of the exoskeleton (epicuticle, exocuticle) are formed before

the old cuticle is shed, ensuring moult-cycle round protection. This cuticula not only enrobes

the entire shrimp’s external body (including eyes and gills), the foregut and hindgut are also

completely covered by a non-permeable cuticle layer (Young, 1959; Thuong et al., 2016b). In

contrast, no cuticle is present to protect the midgut. Instead, a non-cellular, slimy matrix of

chitin and proteins called the peritrophic membrane, ensures local microbial security

(Hegedus et al., 2009). It was experimentally demonstrated that no particles with a diameter

of over 10 nm were able to pass the peritrophic membrane (Martin et al., 2006). The genital

orifices too possess cuticular protection. In open-thelycum species, the female thelycum is a

modification of the chitinous sternal plates of the last two pereopoda. Reception of

spermatophores only occurs during inter- or pre-moult, when the cuticula is at the height of

its protective potential. In so-called closed species, the thelycum is covered by these sternal

plates. The male petasma, responsible for depositing the spermatophores, are also a

modification of the abdominal appendages which are, again, covered by a cuticle (Bailey-


33

Brock and Moss, 1992). It is worth noting that differences in the structure and composition of

the exoskeleton or cuticle exist, depending on its localisation and thus, does not provide

equivalent resistance to noxes. To illustrate, the entire surface of the gills is covered in cuticle,

however uncalcified, and at the height of the lamellae (where oxygen exchange takes place)

the layer is only one µm thick (Bell and Lightner, 1988; Dall et al., 1990).

As discussed above, all potential entry portals are protected by first line defense mechanisms.

To bypass these hurdles, certain conditions must be met for pathogens to succeed in entering

the shrimp and reaching the primary replication sites. Conditions or occurrences that

compromise these barriers can generally be termed risk factors. The identification of risk

factors is often very useful information for applying management measures to prevent

diseases on farm level. Also, it can sometimes lead to the identification of the entry portal.

Thus, good pathogenesis research should never neglect possible risk factors. These conditions

or occurrences can be identified through statistics driven epidemiology research, empirical

research, or sometimes just observational data from shrimp farmers. However, they should

always be confirmed with experiments in a controlled environment to avoid accepting

confounded risk factors as true risk factors.

Epicuticular damage facilitates the entry and nesting of chitinase producing pathogens on the

exocuticle and cause lesions, known as shell disease (Sindermann and Center, 1989).

Conditions in which cuticle damage occur more often, e.g., higher stocking density and

aggression or inferior feed or water quality, Vibrio blooms, directly or indirectly leading to a

weaker cuticula can thus be considered potential risk factors for shell disease. Other possible

risk factors include other (pathogenic or non-pathogenic) organisms, toxins or administration

of antimicrobials, as is often the case with fungal infections such as Fusarium which likes to

act as an opportunistic secondary pathogen. Risk factors can also include conditions leading

to a higher infection pressure or inhibition of the cellular and molecular immunity. Due to the

broad WSSV host range, a lot of risk factors are related to the presence of non-symptomatic

virus carriers such as holes in the fences of farms and nets filtering pumped water or direct

usage of unprocessed seawater (see Tendencia et al. (2011) for a list of articles). Proximity to

the sea (Mohan et al., 2008), use of commercial and fresh feed (Chou et al., 1998; Corsin et

al., 2001), and sharing of pond water (Tendencia et al., 2011) are also risk factors, however


34

rather related to WSSV getting in the pond water than inside the shrimp. Experimentally, it

has been shown that introducing WSSV particles into the water (Chou et al., 1998;

Supamattaya et al., 1998) or cohabitation with infected conspecifics through direct and/or

indirect contact (Flegel et al., 1997; Tuyen et al., 2014) can indeed lead to WSSV-transmission.

Increased stocking density is also linked to WSSV-outbreaks (Tendencia et al., 2011) and can

be interpreted in two ways: more shrimp per m³ water means more infectable and infected

shrimp, which in turn, leads to higher virus titers in the pond water and thus to a higher

infection pressure. High stocking density also leads to increased cannibalism (eating of

infected tissue), aggressiveness (cuticle damage, stress) and increased need to establish social

dominance (higher urination frequency) (Breithaupt and Atema, 2000; Katoh et al., 2008).

Corteel et al. (2009) performed an interesting series of experiments with the goal of assessing

the moulting stage as a possible risk factor for WSSV. No influence in shrimp susceptibility

and mortality was observed when WSSV was injected in shrimp residing in different moulting

stages, meaning no intrinsic variations exist in sensitivity towards the virus over the different

stages. This result indicated that although immunity parameters can vary according to

moulting stage (Liu et al., 2004), the shrimp innate immune response towards this particular

virus remains constant throughout the different stages of the moulting cycle. A different story

emerged when the waterborne route was investigated in relation to the moulting stage. In an

attempt to rule out unwanted cuticle damage caused by the hard tank walls as a confounding

factor, shrimp were immersed in plastic bags containing WSSV. Post-moult shrimp (stages A

and B) were found to be more frequently WSSV-infected than inter- and pre-moult animals

(stages C, D1 and D2) during the immersion challenge in polyethylene bags. When one

pleopod was cut off, even more shrimp in post-moult stages were infected, whereas shrimp

in C, D1, and D2 stages remained uninfected. These series of experiments indicate that during

moulting, first line defenses of the entry portals are compromised rather than the shrimp

innate immunity. Cuticle damage proved to be of some effect, but only if the moulting stage

had already inflicted its effects. A separate study investigating the role of moulting in WSSV-

infection, found 0% mortality when WSSV-immersion-challenge occurred during pre-moult

stages, 53.3% during ecdysis and 26.7% during post-moult (Thuong et al., 2016a).


35

Another potential risk factor that warranted deeper investigation was the effect of salinity

and more precise the transition from one salinity to another. Observations have been made

by shrimp farmers, that WSSV-outbreaks could often be linked to heavy rain falls and after

pond water was changed. Large amounts of freshwater could cause a sudden salinity drop in

open-air ponds. Water mismanagement could hypothetically result in similar consequences.

Experiments mimicking these acute salinity changes indeed found significant effects on the

WSSV-infection rate. Shrimp acclimatised at 35 g l-1 were transferred to lower salinities while

exposed to WSSV. When salinity was reduced to 10 g l-1 and lower, the animals became

infected with WSSV (Thuong et al., 2016a). Reduction of immunity parameters have also been

observed during salinity shock, as well as reactivation in latent WSSV-infected shrimp (Liu et

al., 2006). The following questions can be risen: does a salinity shock make shrimp more

susceptible to WSSV-infection because the immune system is put under stress, or because

the first lines of defense are compromised or maybe a combination of both?

As mentioned before, the exact entry portal is a crucial key stone supporting the pathogenesis

theory. Since the discovery of the virus, a lot of ongoing debate around this subject exists. In

the quest to find how shrimp were getting infected with the, then newly discovered WSSV,

several experimental studies found that feeding WSSV-containing shrimp or feed, effectively

resulted in transmission of the virus (Chang et al., 1996; Wang et al., 1998; Rajendran et al.,

1999). These and subsequent studies led to the conclusion that shrimp were perorally

infected with WSSV and thus, the entry portal was somewhere along the digestive tract.

Overlooking that the tubular epithelial cells of the digestive system are refractive for WSSV-

infection, the presence of a chitinous protective barrier layer and an intact microbiome, the

gut as primary entry portal became the leading consensus. Another way scientists managed

to experimentally infect shrimp (beside intramuscularly) was by water immersion (Chou et

al., 1995; Kanchanaphum et al., 1998; Witteveldt et al., 2004, 2006; Corteel et al., 2009). It is

important to note that these studies were not performed under the most strictly controlled

conditions, even viral titrations of the inoculates were often not performed. Efficiency of

these kind of infections was low and sometimes difficult to reproduce (Perez et al., 2005;

Laramore, 2007; Corteel et al., 2009). Transovarial infection has been described as well: WSSV

can be found in testes, ovaries, follicular cells, oogonia and oocytes (Lo et al., 1997). However,

it is worth noting that per ovum infections are always retraceable to previous generation(s)


36

of shrimp, where the pathogen first succeeded to infiltrate via one of the six aforementioned

primary entry portals.

Previous experiments have indubitably shown that moulting stage influences WSSV-entry.

During the moulting stage, endogenous chitinases are released that could possibly alter the

protective capabilities of the cuticula or peritrophic membrane enough to allow for WSSV-

entry (Tan et al., 2000; Proespraiwong et al., 2010). This would explain, why feeding of WSSV-

contaminated feed to shrimp results, although with low efficiency, in WSSV-positive shrimp

(most of the peroral inoculation studies did not account for moulting stage). In insects, it was

shown that inhibition of this peritrophic membrane, leads to a higher susceptibility to several

pathogens (Wang and Granados, 2000; Rao et al., 2004) and in shrimp, Vibrio

parahaemolyticus, and Vibrio harveyi were shown to use bacterial chitinases to destroy and

penetrate the peritrophic membrane in order to colonise the midgut, and causing syndromes

like acute hepatopancreatic necrosis (Martin et al., 2004; Tran et al., 2013). Endochitinases

would weaken the shell and make shrimp more vulnerable for wounds or small fissures.

Moreover, the midgut was previously suggested as possible entry portal for WSSV by several

authors (Di Leonardo et al., 2005; Arts et al., 2007). Thuong et al. (2016b) investigated

whether a compromised peritrophic membrane could indeed facilitate WSSV-entry at the

height of the midgut. They used saltwater to gently flush out the peritrophic membrane

before performing titration experiments (Figure 2). However, the authors did not retrieve

data supporting the claim that removal of this structure significantly facilitates WSSV-entry

and that the midgut is an entry portal for WSSV. The tubular epithelial cells of the midgut are

not receptive to WSSV and together with the basal membrane, they provide an effective

barrier against WSSV-colonisation of the shrimp, even when the peritrophic membrane is

compromised. The method of Thuong et al. (2016b) could however be useful to test the entry

of several other shrimp pathogens via the midgut.

Thus, despite all these efforts, the site where WSSV-entry occurs, remains enigmatic.

Therefore, other potential entry portals should be investigated. The antennal gland and

integumental glands surfaces: those are the only two structures which are connected to the

outside world and of which the lumen is not covered by an exoskeleton or chitinous layer

(Young, 1959; Juberthie-Jupeau and Crouau, 1977). The tegumental glands are clusters of


37

cells, located underneath the epidermal cell layer. The secrete of this gland is collected by a

central duct, leading towards pores in the shrimp’s outer surface (Felgenhaur, 1991). To which

extent the integumental glands play a role in shrimp infectious diseases and how or if the

gland is protected, remains unclear and merits future research. The antennal gland appears

to be a prime candidate for WSSV-entry site. First and foremost, WSSV is able to infect

antennal gland cells (Lo, 1996). Furthermore, as discussed above, it has been shown that

salinity changes increase WSSV-susceptibility (Thuong et al., 2016). Since the antennal gland

plays a lead role in osmoregulation and hemolymph filtration, and some parts during

moulting, it is possible that the antennal gland is indeed involved in WSSV-entry. Although

the access to the antennal gland is controlled by the chitinous nephropore, the rest of the

organ’s lumen is entirely devoid of cuticular lining and thus vulnerable to pathogen invasion

(Young, 1959). Because of these arguments, the antennal gland of the P. vannamei, will be

the subject of investigation in this thesis.

1.3. GENERAL MORPHOLOGY OF THE PENAEID SHRIMP

Figure 6. General external morphology of the Penaeus vannamei A: rostrum B: carapax A+B: cephalothorax, C:

body length, D: abdomen, E: antennulae, F: stylocerite, G: scaphocerite (in abducted position), H: endopodite of

the antenna, I: flagellum of the antenna, J: maxillipodae, K: pereiopodae, L: pleopodae, M: uropoda, N: exopodite

of the uropodite, O: endopodite, P: telson, Q: dorsal crest (Chan, 1998)


38

To situate the morphology and anatomy of the antennal gland, an overview of the general

anatomy of the penaeid shrimp will be given first. The external morphology of the shrimp is

shown in Figure 6. Like in all crustaceans, the shrimp’s body can be roughly subdivided in two

major sections: (i) the cephalothorax, which contains almost all organs, and (ii) the abdomen,

which consists mainly of muscle tissue. Two pairs of antennae are present on the stylocerites

(first segments of the cephalothorax). Additionally, one pair of larger antennae (also called

the second antennae or terminal flagellae) can be found on the pediculum (appendix of the

second segment). Thus, on both sides, the shrimp possesses three antennae on both sides (six

in total) (Young, 1959). On the base of the antennal pediculum, a chitinous structure, the

nephropore, is present. This pore contains the orifice (opening) to the antennal gland, which

is the excretory gland of the shrimp. The nephropore is located at the height of the proximal

segment of the antenna, which is referred to as the coxipodite or coxicerite (Figure 7). The

coxipodite is the first segment of the pediculum and forms a rigid articulation with the

basipodite or basicerite distally and the cephalothorax proximally. Protopodite is a term

sometimes used to denote these first two segments (coxipodite plus basipodite). On the distal

side of the basipodite, both the endopodite (medial) and the exopodite (lateral) are

implanted. The antennal exopodite is better known as the scaphocerite or the antenna shell.

The endopodite can in its turn be subdivided from proximal to caudal in the ischiocerite,

merocerite and carpocerite. It is on this endopodite that the annularly segmented terminal

flagellum or antenna is implanted. These antennae are the etymological base for the antennal

gland (Young, 1959; Stachowitsch, 1992).


39

Figure 7. The appendages of the second segment on the cephalothorax of the Penaeus vannamei: the

second pair of antennas or antennae Left: ventral view and right lateral view. A: coxipodite or

coxicerite B: basipodite or basicerite C: ischiocerite, D: merocerite, E: carpocerite, F: flagellum or

(second) antenna, G scaphocerite or exopodite of the antenna, O: nephropore, A+B+C+D+E:

pediculum of the antenna, A+B: protopodite of the antenna, C+D+E: endopodite of the antenna.

1.4. THE ANTENNAL GLAND

1.4.1. Introduction

In contrast to mammals, crustaceans seem to have multiple organs responsible for excretion,

osmoregulation, and ion-metabolism: the antennal gland (also known as the green gland,

renal gland, and antennary gland), the maxillary gland and the gills. The latter are not only

known for their respiratory function, where oxygen and carbon dioxide are exchanged

between the water and the hemolymph, but also for ammonium regulation, osmoregulation,

calcium uptake, and acid/base regulation (Foster and Howse, 1978; Taylor and Taylor, 1992;

McMahon, 1995; Bauer, 1999; Ahearn et al., 2004; Freire et al., 2008). Except for the

respiratory function, the antennal gland and the maxillary gland perform grosso modo the

same functions as the gills. However, the antennal gland and the maxillary gland do not

appear together in the same species (Potts and Parry, 1965). They both have a similar

blueprint, cytology, and function, but when the organ exits the animal at the base of the

antennal coxipodite, it is called the antennal gland and when it opens at the maxillae, it is

called the maxillary gland. As mentioned above, shrimp possesses antennal glands. The

similarities between the excretory organs of Decapoda and mammals go as far as the


40

description by Maluf (1941) of some kind of a kidney stone in the coelomosac of the crayfish

Cambarus clarkii.

The crustacean excretory organ is a bilaterally built, segmented coelomoduct organ, and can

generally be subdivided into three main parts: (i) a coelomosac, (ii) an efferent duct, and (iii)

a terminal duct, leading to a nephropore (Young, 1959; Icely and Nott, 1979; Freire et al.,

2008). The coelomosac is a terminal ‘bag’ of mesodermal origin and is responsible for primary

ultrafiltration of the hemolymph, it is thought to be a remnant of the coelom. Next to the

coelomosac is the efferent duct. In decapods, this has evolved into an anastomosing network

of tubuli, called the labyrinth (Peterson and Loizzi, 1973). The terminal duct sometimes has a

urinary bladder, but always leads to the exit pore, namely the nephropore. These different

segments together form a single nephron-like structure that in many aspects resembles the

different parts of a vertebrate kidney (Beams et al., 1956; Johnson, 1980; Roldan and Shivers,

1987; Fuller et al., 1989; Behnke et al., 1990; Fingerman, 1991; Nakamura and Nishigaki, 1991;

Felgenhaur, 1992; Nakamura and Nakashima, 1992; Lin et al., 2000; Vogt, 2002; Xiaoyun et

al., 2003; Wheatly et al., 2004; Al-Mohsen, 2009; Buranajitpirom et al., 2010; Xugang et al.,

2013). The nephropore is located near the coxipodite of the antennae; hence the name

'antennal gland' (Young, 1959; Nakamura and Nishigaki, 1991). Both the coelomosac and the

efferent duct are thought to be of mesodermal origin, whereas the terminal duct is

ectodermal tissue (Anderson, 1973; Fingerman, 1991; Freire et al., 2008). Many hiatuses still

exist concerning the different structures of the nephrocomplex. Moreover, the vast body of

today’s knowledge is extrapolated from other crustaceans, mainly crayfish and lobster.

Below, the (ultra)structure, morphology, and anatomy of the different parts of the antennae

gland of Decapoda will be described and the functional role will be addressed when possible.

1.4.2. Functions of the antennal gland

The life cycle of the euryhaline species P. vannamei demonstrates the capacity of the marine

animal to tolerate an extreme range of salinity from brackish (0.5 g/L) to high marine salinities

(60 g/L), when given enough time to gradually acclimatise (Ayaz et al., 2015). The salinity at

which optimal growth is observed, is around its isosmotic point (ca. 25‰) (Castille and

Lawrence, 1981; Ponce-Palafox et al., 1997; Jannathulla et al., 2019). However, more factors

are to be considered when selecting for optimal salinities like response to stress and disease.


41

It can also be of no surprise that the overall optimal salinity at which shrimp should be reared,

varies with age, given their life cycle in the wild (Li et al., 2017). Crustaceans are supposed to

be osmocomformers, however, osmoconformers are stenohaline, while P. vannamei is an

extremely effective euryhaline species (Ayaz et al., 2015; Rivera-Ingraham and Lignot, 2017).

Furthermore, Lin et al., (2000) showed that P. vannamei responds to salinity changes by

expelling or ceasing urine excretion. This would make P. vannamei an osmoregulator. Perhaps

penaeid shrimp are both, depending on present conditions. Future research will have to be

performed to elaborate the exact mechanism of osmoregulation in Penaeid shrimp.

Maintaining body volume and hemolymph osmolarity are the most evident functions of the

antennal gland. When shrimp are subjected to sudden drops in salinity, more urine is

produced to keep the shrimp and its cells from swelling and bursting. Vice versa, when shrimp

are exposed to a high salinity environment, the production of urine is halted to prevent

shrimp from dehydrating (Lin et al., 2000). As a reaction to changes in salinity, but also during

other circumstances, the shrimp’s antennal gland is known to regulate body ion levels like

magnesium, calcium, potassium, sulphate, copper, iron, and sodium (Dall and Smith, 1981;

Cheng and Liao, 1986; Vogt and Quinitio, 1994; Lin et al., 2000). Na+/K+-ATPase activity in the

antennal gland, is related to the salinity of the water in which shrimp are reared

(Buranajitpirom et al., 2010). Another saline sensitive activity in the antennal gland, is that of

the carbonic anhydrase metalloenzyme. Carbonic anhydrase catalyses the interconversion

between carbodioxide and bicarbonate in the presence of water. It is believed to rapidly

supply counter ions for transport processes involved in ion regulation, like NaCl uptake

against a chemical gradient (Henry et al., 2006; Pongsomboon et al., 2009). Carbo anhydrase

has been reported in many invertebrates and is ascribed to play an active role in the

acid/base-regulation of Penaeus vannamei as well (Liu et al., 2015).

When shrimp are exposed to heavy metals, the antennal gland, along with several other

organs acts as a chelator and excretes these toxic components. Mercury, lead, chromium,

nickel, zinc, cadmium, and manganese are described to be sequestered and secreted via the

urine when animals were challenged with these toxic elements. Thus, the antennal gland

plays an important role in the detoxification of shrimp (Doughtie and Rao, 1984; Vogt and

Quinitio, 1994; Páez-Osuna and Tron-Mayen, 1996; Xugang et al., 2013). The presence of fixed


42

phagocytes hint at a possible role in the immune system, either by eliminating threats or

cleaning remaining fragments (Johnson, 1987; Kondo et al., 2012).

Like many other animals, shrimp fight for feed and to establish social relations and/or

dominance. In Macrobrachium species and lobsters, social dominance involves the excretion

of hormones via the urine (Breithaupt and Atema, 2000; Katoh et al., 2008; Al-Mohsen, 2009).

When shrimp are getting ready to mate, the egg laying hormone was found in the shrimp’s

antennal gland, potentially regulating the hemolymph hormone level, or maybe acting as a

signalling peptide to other shrimp (Liu et al., 2006). In Macrobrachium rosenbergii, the male

to female attraction stimulating hormone “N-acetylglucosamine-1,5-lactone” was found to

be expressed in its excretory organ (Bose et al., 2017) and in Sagmariasus verreauxi,

expression of genes with pheromone function were identified in this organ (Chandler et al.,

2016). Together with the discovery of the “trapped in endoderm receptor 1” (suggested to be

crucial in courtship and mating behaviour in Drosophila melanogaster) in the antennal gland

of P. monodon, these studies strongly hint at the antennal gland as a pheromone releasing

organ in shrimp (Viet Nguyen et al., 2020).

The antennal gland also plays a role in the moulting process and cycle. The moulting hormone

ecdysone from the Y-organ is metabolised by the antennal gland and its metabolites can be

excreted by the same route. Consequently, the antennal gland can regulate the balance

between the moult inhibiting hormone and moulting hormone ecdysone. The latter

determine the subsequent moulting stages (Chang, 1985; Mykles, 2011). During stage D1

(early pre-moult), the moulting hormone ecdysone reaches a peak, signalling the cells to

prepare for ecdysis. When the shrimp reaches pre-moult, these hormone levels drop

drastically (Corteel and Nauwynck, 2010). Finally, by regulating the shrimp’s calcium balance,

the nephrocomplex can regulate the hardening of the new, still soft shell by mineralisation.

Also, calcium plays an important role as an intracellular messenger in the hormonal actions

(Ahearn et al., 2004; Wheatly et al., 2004). Water uptake immediately before ecdysis is of

great importance, since it allows animals to break out of the shell and the amount of water

will determine the final volume of the crustacean when the cuticle hardens in the next hours

or days. It is believed that they take up water by drinking (Mykles, 1980; Dall et al., 1990).


43

1.4.3. Coelomosac

The coelomosac is the place where the ultrafiltration takes place (Behnke et al., 1990;

Nakamura and Nishigaki, 1991; Felgenhaur, 1992). The ultrastructure of the coelomosac

seems to be constant across the different Crustacea. Its functional cells (podocytes) form a

monolayer epithelium and highly resemble the podocytes found in the Bowman capsule of

mammals. Intercellular channels between the podocytes can be found and adjacent cells are

connected by desmosome-like junctions. Cytoplasmatic cytopodia connect the podocytes

with the basement membrane (Schmidt-Nielsen et al., 1968; Ueno et al., 1997; Khodabandeh

et al., 2005a; Al-Mohsen, 2009; Xugang et al., 2013). On top of the basement membrane, a

slit diaphragm was found in crayfish (Fig 5). This diaphragm acts as a secondary filtration

barrier, blocking macromolecules which manage to pass the basement membrane. These

blocked macromolecules form a lamina densa between the slit diaphragm and the basement

membrane (Schaffner and Rodewald, 1978; Vogt, 2002). The podocytes contain organelles

which indicate active producing cells (golgi-apparatus, mitochondria, rough endoplasmatic

reticulum, vesicles, vacuoles, ribosomes, and a big nucleus with clear nucleoli, Fig 5).

How the coelomosac is fed with hemolymph is not completely clear. Usually, the coelomosac

is surrounded by haemolymph sinuses fed by arteries (Panikkar, 1941; Schmidt-Nielsen et al.,

1968; Schaffner and Rodewald, 1978; Tikær Andersen and Baatrup, 1988; Ueno and Inoue,

1996; Vogt, 2002; Khodabandeh et al., 2005a; Al-Mohsen, 2009; Tsai and Lin, 2014). Other

researchers mention blood vessels in direct contact with the basal membrane of the

coelomosac cells (Holliday and Miller, 1984; Xiaoyun et al., 2003). Still others report that the

coelomosac of the P. japonicus bathes freely in the haemocoel (Nakamura and Nishigaki,

1991). Eventually, fluid will pass the basement membrane and the slit diaphragm to become

ultrafiltrate. Next, when the ultrafiltrate flows through the intercellular channels between the

podocytes, these cells modify the fluid by pinocytosis (Peterson and Loizzi, 1974; Ueno and

Inoue, 1996). Likewise, substances from the coelomosac lumen are endocytosed (Kirschner

and Wagner, 1965). Doing so, podocytes can capture and sometimes metabolise ions

(Schmidt-Nielsen et al., 1968; Holliday and Miller, 1984), metallic particles for e.g.,

detoxification (Doughtie and Rao, 1984; Roldan and Shivers, 1987), proteins, and other

(macro)molecules (Ueno and Inoue, 1996; Ueno et al., 1997). Both endocytic and pinocytotic

vesicles will fuse together to form dense bodies, which are excreted by a process called


44

blebbing (Fig 5) (Ueno and Inoue, 1996). Once in the coelomosac lumen, the dense bodies are

called formed bodies (Riegel, 1966a; Khodabandeh et al., 2005b). In crayfish, these formed

bodies are studied best. It is believed that when they are in the coelomosac lumen, they still

contain large, undigested macromolecules (Riegel, 1966b, a). Finally, at the basal side of the

cytopodia, Na+/K+ ATPases are present in crabs, as well as H+-ATPases, Na+/K+/2Cl--

cotransporters and Na+/H+ exchangers in the apical side of the podocytes (Tsai and Lin, 2014).

Khodabandeh et al. (2005) however, could not find Na+/K+-ATPases in the lobster coelomosac.

In Penaeus monodon, Buranajitpirom et al. (2010) found Na+/K+-ATPase immunoreactivity in

the coelomosac. When shrimp were challenged to different salinities, the enzyme activity

inside the antennal gland was altered in adaptation to these conditions.

Figure 8. Ultrastructure of the coelomosac Pc: podocyte, bM basal membrane, H: hemolymph, CL: coelomosac

lumen, CP: cytopodia, ICK: intercellular cannulas, SD: slot diaphragm, V: phagocytotic vesicles, G: golgi device, E:

endosomes, N: nuclei, n: nucleoli, DL: dens body, B: blebbing, GL: formed body, Hc: hemocyte. The red arrow

indicates the flow of fluid from the hemolymph to the coelomosac lumen. The dense bodies are excreted by the

cells through a process called 'blebbing'. The whole is squeezed off (*) and is now part of the urine.


45

1.4.4. Labyrinth

Eventually, the collected filtrate inside the coelomosac will flow through to the labyrinth. The

labyrinth is made up of anastomosing tubuli, bathing in the hemolymph and, in shrimp,

surrounding the coelomosac (Nakamura and Nishigaki, 1991; Nakamura and Nakashima,

1992; Xiaoyun et al., 2003). In contrast to crayfish, lobster and Macrobrachium species, only

one type of labyrinth cell exists, a single-cell-layer thick cuboidal to high columnar epithelium,

with a microvilli brush border and a basally located nucleus (Xiaoyun et al., 2003; Xugang et

al., 2013). This cytomorphology strongly suggests absorption, reabsorption and/or other

modifications of the filtrate. Indeed, it is involved in reabsorption of large molecules, fluid,

and ions. Waste elimination and regulation of osmotic pressure are also attributed to the

labyrinth tubuli (Xiaoyun et al., 2003). Like in the lobster (Khodabandeh et al., 2005), Na+/K+-

channels were found inside certain cells of the penaeid labyrinth (Buranajitpirom et al., 2010).

Formed bodies coming from the coelomosac start to undergo some changes once arrived in

the lumen of the labyrinth (where a lower pH exists). There, pH-dependent proteases are

activated, digesting the contents of the formed bodies. Subsequently they attract water and

burst. Together with hydrolysis processes it is believed that in crayfish, this process creates a

concentration gradient between the hemolymph and the lumen of the labyrinth effectively

aiding in the osmoregulation (Riegel, 1966a, b, 1970a, b). Whether this is the case in shrimp

remains to be examined.

1.4.5. Terminal duct

Following the labyrinth, the filtrate arrives in the terminal ductus. In Penaeid shrimp, this

ductus has evolved in a urinary bladder containing 2 cell layers. One basal layer, topped by a

flattened to columnar epithelium with short microvilli. This conformation gives the structure

a better flexibility as its function is most likely to retain urine before evacuation (mictio)

although reabsorption has been described too. Finally, the urinary bladder will slim down to

a short duct, leading to the nephropore (Vogt, 2002; Xiaoyun et al., 2003).

1.4.6. Overall morphology

Generally speaking, the blueprint of the bilaterally built antennal gland is very species specific.

In Penaeid shrimp, the tubuli of the labyrinth are organised around the central coelomosac.

The labyrinth flows into a urinary bladder and finally via a short ductus to the exit pore (the


46

nephropore), which is connected to the outer world (Xiaoyun et al., 2003). Young (1959) who

did a very thorough investigation of penaeid morphology, described colourations above the

supra-esophageal ganglion after dye injection in the nephropores, hinting at an unknown part

of the antennal gland. Several other studies reported small tubules spread throughout the

hemocoel. These tubuli even make functional connections with the lymphoid organ

(Duangsuwan et al., 2008; Rusaini and Owens, 2010). Finally, a Japanese team attempted to

reconstruct the anatomy and morphology of the antennal gland in Penaeus japonicus, using

a clay model based on histological sections. They described the coelomosac to be made of

three sacs: (i) a median sac centrally containing the labyrinth, (ii) the transversal sac, sprouting

from the medial side of the median sac, connecting both halves of the antennal gland just

caudally from the brains, (iii) a median lobe with a short epigastric lobe. This median lobe

originates from the caudal side of the transversal sac and will cover the stomach rostrally.

According to the authors, the median lobe is the place of ultrafiltration (Nakamura and

Nishigaki, 1991; Nakamura and Nakashima, 1992). It is worth noting that functionally, this

study does not comply with all other functional morphological studies (Lin et al., 2000;

Xiaoyun et al., 2003; Buranajitpirom et al., 2010; Xugang et al., 2013).


47

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Xugang, H., Guangfu, H., Guangtao, L., 2013. The effect of lower salinity on microstructure of antennary gland of Litopenaeus vannamei. Journal of Food, Agriculture & Environment 11, 782-785. Young, J., 1959. Morphology of The White Shrimp Penaeus setiferus (Linnaeus 175), Fishery Bulletin 145. The Fish and Wildlife Service, United States of America, p. 173. Zhan, W.-B., Wang, Y.-H., Fryer, J.L., Yu, K.-K., Fukuda, H., Meng, Q.-X., 1998. White Spot Syndrome Virus Infection of Cultured Shrimp in China. Journal of Aquatic Animal Health 10, 405-410. Zhu, F., Twan, W.-H., Tseng, L.-C., Peng, S.-H., Hwang, J.-S., 2019. First detection of white spot syndrome virus (WSSV) in the mud shrimp Austinogebia edulis in Taiwan. Scientific Reports 9, 18572. Zwart, M.P., Dieu, B.T.M., Hemerik, L., Vlak, J.M., 2010. Evolutionary Trajectory of White Spot Syndrome Virus (WSSV) Genome Shrinkage during Spread in Asia. PLOS ONE 5, e13400.

Chapter 2

Aims and Outline of the thesis

G.M.A. De Gryse



Chapter 2 Aims

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Ever since its first emergence, white spot syndrome virus (WSSV) has been causing serious

havoc in shrimp aquaculture worldwide (Sanchez-Paz, 2010; Stentiford et al., 2012; Patil et

al., 2021). This OIE-listed notifiable disease causes massive losses due to its high mortality and

morbidity, both up to 100% (Nunan et al., 1998).

A missing piece in the pathogenesis puzzle is the entry portal and the primary replication site.

In Chapter 3 the morphology of the antennal gland was investigated in order to find out if this

excretory organ is a suitable candidate for pathogen entry by a 3D-reconstruction of the organ

by stacking haematoxylin-eosin-stained sections of the shrimp cephalothorax.

In Chapter 4, a micro magnetic resonance imaging (µMRI) technique was used to confirm the

morphology of the excretory organ obtained in Chapter 3 in vivo and to determine its

involvement in the moulting cycle.

A pathogen must breach primary host defences before it can reach its primary replication site

and invade its host towards secondary replication sites. Thus, in Chapter 5 the biodynamics

and defensive prowess of the nephropore in protecting the access to the antennal gland were

studied and tested using scanning electron microscopy and ex vivo modelling. The intrinsic

risk of urination was also investigated.

Finally, the true capacity of pathogens to enter, infect and disease shrimp was checked in

Chapter 6. Both viral (WSSV) and bacterial (Vibrio) pathogens were inoculated through the

nephropore into the bladder. Finally, a salinity shock was used as a risk factor to increase the

frequency with which shrimp urinate to facilitate and demonstrate natural WSSV infection.

Chapter 2 Aims

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References Patil, P.K., Geetha, R., Ravisankar, T., Avunje, S., Solanki, H.G., Abraham, T.J., Vinoth, S.P., Jithendran, K.P., Alavandi, S.V., Vijayan, K.K., 2021. Economic loss due to diseases in Indian shrimp farming with special reference to Enterocytozoon hepatopenaei (EHP) and white spot syndrome virus (WSSV). Aquaculture 533, 736231. Sanchez-Paz, A., 2010. White spot syndrome virus: an overview on an emergent concern. Veterinary Research 41. Stentiford, G.D., Neil, D.M., Peeler, E.J., Shields, J.D., Small, H.J., Flegel, T.W., Vlak, J.M., Jones, B., Morado, F., Moss, S., Lotz, J., Bartholomay, L., Behringer, D.C., Hauton, C., Lightner, D.V., 2012. Disease will limit future food supply from the global crustacean fishery and aquaculture sectors. Journal of Invertebrate Pathology 110, 141-157.

Chapter 3

Anatomical and morphological

exploration of the antennal gland

G.M.A. De Gryse



Adapted from: De Gryse, G.M.A., Khuong, T.V., Descamps, B., Van Den Broeck, W., Vanhove, C., Cornillie, P., Sorgeloos, P., Bossier, P., Nauwynck, H.J., 2020. The shrimp nephrocomplex serves as a major portal of pathogen entry and is involved in the molting process.

Proceedings of the National Academy of Scienses U S A 117, 28374-28383.

Chapter 3 Morphology of the antennal gland

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3.1. ABSTRACT

White spot syndrome virus (WSSV) wreaks havoc on shrimp industry. As the main portal of

entry remains unclear, infectious diseases such as WSSV remain a difficult issue to tackle in

the field. Because the cuticle provides a strong barrier against invading pathogens, regions

lacking cuticular lining, such as the shrimp’s excretory organ “the antennal gland” are major

candidate entry portals. The antennal gland is up till now underexplored in penaeid shrimp

and especially its anatomy and morphology. In this chapter, histological techniques are

applied together with specialised 3D-software to reconstruct a 3D-model of the antennal

gland and its surrounding and associated organs. We demonstrated that the antennal gland

resembles the blueprint of the mammal kidney, with a newly described tubular network to

the rostral side of the animal and a complex of bladders reaching as far as the middle of the

hepatopancreas. The muscles associated with this organ were also reconstructed and studied

using the same techniques. Several observations could back the hypothesis of the antennal

gland as a major pathogen entry portal and links were made to the moulting cycle, which is

in turn also linked to WSSV-infection. Finally, based on all data, a new name was proposed for

the antennal gland, i.e. “The nephrocomplex”.

Key words: Penaeus vannamei, WSSV, antennal gland, anatomy, morphology


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3.2. INTRODUCTION

Shortly after the emergence of the devastating white spot syndrome virus (WSSV) in shrimp

aquaculture, it was thought that WSSV entered its host upon ingestion of viral particles.

Success with feeding contaminated shrimp carcasses led to a long-standing consensus on the

per oral route for WSSV-entry, although a direct proof was not demonstrated (Chang et al.,

1996; Escobedo-Bonilla et al., 2008). The gut is made up of three regions: the foregut, the

midgut, and the hindgut. Both fore- and hindgut are covered with an impenetrable layer of

cuticula (Bell and Lightner, 1988; Corteel et al., 2009), while the midgut is protected from

pathogens by a peritrophic membrane (Thuong et al., 2016). Thus, not surprisingly, the

efficiency with which shrimp can be infected per os is extremely low and the gut as WSSV

entry portal is still controversial (Domínguez-Borbor et al., 2019; Thuong et al., 2016; Wang

et al., 1998). Therefore, the search for new possible organs which allow WSSV to enter the

animal remains crucial in pathogenesis-oriented research.

The crustacean antennal gland has been the subject of research for many years. While this

organ has been studied extensively in crab, lobster and crawfish species, little work has been

conducted concerning the penaeid shrimp antennal gland (see chapter 1, introduction). As

this organ is not covered with cuticula and is connected with the outer world via the

nephropore, it is one of the main candidate portals of entry for pathogens. Significant parts

of the antennal gland are of mesodermal origin, which are highly susceptible for WSSV

(Anderson, 1973; Escobedo-Bonilla et al., 2008). It is of no surprise that WSSV-replication

inside the antennal gland has been described on many occasions (Chang et al., 1996;

Rodríguez et al., 2009). Moreover, several pathogens have already been found inside the

antennal gland of other crustaceans. Thrupp et al. (2013), for example, found a parasitic

infection in the antennal gland of juvenile crabs. In the same manner, Pina et al. (2007)

isolated Cercaria sevillana from the green crab’s antennal gland, whereas other researchers

found a herpesvirus-like particle through electron microscopical examination of the antennal

gland in King crabs (Ryazanova et al., 2015). The antennal gland, together with the gills, are

responsible for the excretion of metabolites like moulting hormones and the regulation of the

body volume (Buranajitpirom et al., 2010; Lin et al., 2000). Induced salinity shock has been


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proven to be linked to WSSV-infection, as is moulting (Corteel et al., 2009; Thuong et al.,

2016).

The antennal gland is bilaterally present and is composed out of a primary filtration unit (the

coelomosac), an efferent secondary filtration module (labyrinth) and a terminal duct leading

to the nephropore, the exit hole of the excretory organ (Buranajitpirom et al., 2010; Freire et

al., 2008; Icely and Nott, 1979; Young, 1959). Based on structure and function, the antennal

gland can be seen as the homologous counterpart of the mammalian excretory system. The

coelomosac is located in the front third of the cephalothorax at the height of the antennal

appendix, lined by podocytes and responsible for ultrafiltration of the hemolymph. This

filtrate can be modified by pinocytosis and endocytosis of ions and other (macro-)molecules

as it flows through the coelomosac lumen (Felgenhaur, 1992; Holliday and Miller, 1984;

Xiaoyun et al., 2003). By excretion of formed bodies in the coelomosac of Crayfish, a

concentration gradient can be formed to help attract water from the hemolymph at the

height of the labyrinth (Riegel, 1966a, 1966b, 1970). This labyrinth is where the ultrafiltrate

flows from the coelomosac. It is composed of highly anastomosing tubuli (Peterson and Loizzi,

1974). The columnar pseudo multiple cell layer with observed cytoplasmatic invaginations is

apically equipped with a microvilli brush boarder, indicative for (re)absorption, and thus

further modification of the ultrafiltrate (Buranajitpirom et al., 2010; Xugang et al., 2013).

Finally, the modified ultrafiltrate from the labyrinth, is ventrally collected in a urinary bladder.

This is a modification of the evolutionary terminal duct. The bladder is two cell layers thick,

columnar, or flattened, built for flexibility. It is said to be functioning as a temporal storage

place (Khodabandeh et al., 2005; Vogt, 2002). A short duct finally leads to the nephropore,

through which the urine can be expelled (Young, 1959).

In penaeids, macroscopically, the antennal gland can be described as a bean-shaped beige

organ, located at the base of the antennal appendix (Buranajitpirom et al., 2010). The

conformation and exact anatomical blueprint of the aforementioned structures are subject

of debate and differ among species (see introduction). Moreover, a Japanese clay model

reported different modifications of the coelomosac, limited to rostral of the hepatopancreas

and caudal of the brain, in the form of three coelomosac bladders (Nakamura and Nishigaki,

1991; Nakamura and Nakashima, 1992). Finally, Young (1959) suspects parts of the antennal


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gland to be located above the brain, after injection of coloured fluids in the antennal gland.

In short, the anatomy of the penaeid shrimp is up till now not indefinitely described. In this

chapter, an attempt was made to fulfil this hiatus, as the antennal gland is a prime suspect to

serve as entry portal for WSSV.

3.3. MATERIALS & METHODS

3.3.1. Ex vivo 3D-reconstruction of HE-slices with AMIRA

Animals

Early pre-moult SPF P. vannamei shrimp with mean body weight (MBW) of 9.13 ± 0.92 g were

imported from the Netherlands and reared in the Artemia Reference Center (ARC, Faculty of

Bioscience engineering, Ghent University, Belgium). There, they were screened for moulting

stage C according to Corteel et al. (2012) by inverted light microscopical inspection of the

uropods (tail fan) and subsequently brought to the Laboratory of Virology (Faculty of

Veterinairy Science, Ghent University, Belgium). At the Laboratory of Virology, the shrimp

were allowed to acclimatise for 2 days at 28°C in separate aerated 10 L PVC tanks at a salt

concentration of 35 g/L. After this acclimatisation period and recovery from possible

transport related stress, they were euthanised by slowly adding more and more ice to the

PVC tank until movement stopped. Next, the animals were transported on ice to the

Laboratory for Light and Electron Microscopy, Department of Morphology (Faculty of

Veterinary Science, Ghent University, Belgium).

Sample preparation

The cephalothoraxes were cut from the abdomen without compromising the hemocoel space

to keep internal configuration intact as much as possible. Next, the cephalothorax was fixed

by injection of Bouin fixative, a solution of paraformaldehyde and picric acid (85 mL of 2%

paraformaldehyde and 15 mL of saturated picric acid, pH 7.4) and mobile immersion in the

same solution at 37°C for 12 h. Following an overnight washing step in 70 ethanol, the

cephalothoraxes were dehydrated with ethanol and xylene in a STP 420D Tissue Processor

(Microm, Prosan, Merelbeke, Belgium) and imbedded in paraffin with an EC 350-2 Modular

Tissue Embedding Centre (Microm, Prosan, Merelbeke, Belgium). Ten µm thin slices were


79

made using a HM360 Rotor Microtome with water bath (Microm, Prosan, Merelbeke,

Belgium) and caught on gelatine coated microscope slides. After drying, the sections were

stained with a classic hematoxylin-eosine (HE) staining. Finally, cover slips were mounted with

a drop of mounting fluid (DPX, Sigma-Aldrich, St. Louis, USA).

Image acquisition & data input

The slides were visually evaluated using an Olympus BX50 light microscope (Aartselaar,

Belgium) and scored for intactness of the antennal gland’s related structures and suitability

for 3D-reconstruction. With Cell F-software, serial digital image acquisitions were made with

a 20x magnification lens at standard resolution and saved in .JPEG format. A ‘stacked slices’

format friendly .txt-file was created containing all specific .JPEG file names, with relative

distance to the first image file, together with the pixel size. Using AMIRA v6.0 (FEI Visualization

Sciences Group Europe, Mérignac, France), both .txt-files and digital acquisitions were

uploaded and saved as a ‘stacked slices file’ format.

Image processing & analysis

In AMIRA, using the ‘least square method’ in the ‘align slices module’ subsequent slices were

aligned automatically and later manually adjusted when necessary. Aligning was performed

according to the orientation of adjacent slices or the closest correct slice. Next, slices were

resampled in a ‘mesh file’ format. Because HE-stainings are dominantly red and purple, the

application of a green filter was chosen to achieve a greater contrast using the ‘cast field’

function to create a grey scale image, necessary for the next step of segmentation; by

calculating the difference in greyscales between the voxels, various structures become visible

on 2D slices.

The structures of interest could be traced on a 2D-plane by selecting voxels of interest and

labelling them. Different labels of different colour for different structures of interest were

applied. To select the voxels of interest, the threshold tool, lasso tool, magic wand tool, draw

limit line, brush tool, pick and move tool, and blow tool were used. When stacking the serial

2D-labels, a 3D-image of several structures of interest was created. If adjacent sections did

not contain sufficiently intact structures or when the alignment was off, the interpolation

function was used to calculate labels to uphold continuity of the 3D-image in order to keep


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resolution along the Z-axis. The actual 3D-reconstruction was performed using the

SurfaceGen module (Generate Surfaces) with constrained smoothing and minimal edge

length of 0.4 to smooth and reduce the complexity of the model without losing voxel data. A

final smoothing step was undertaken in de Surfaceview module by normalising the vertex

normal (nodes). To evaluate the antennal gland at cellular level, an Olympus BX50 light

microscope (Aartselaar, Belgium) with several magnifications was used.

3.4. RESULTS

3.4.1. General overview of the antennal gland morphology

Three-dimensional reconstruction of the antennal gland and surrounding organs by

superposition of serial haematoxylin and eosin-stained sections (10 µm) of the cephalothorax

revealed a much more complex and widespread organization of this organ than previously

believed (Fig. 1). The entire antennal gland can be divided into several compartments. The

three main divisions found in other crustaceans (coelomosac, labyrinth and terminal duct with

urinary bladder) could also be found here in penaeid shrimp, but several modifications were

found reaching throughout the entire cephalothorax. The coelomosac and labyrinth are

gathered within a well-defined compact glandular part (CGC). To the rostral side of the animal,

another body of tubuli was observed, named ‘the rostral compartment’ (RC), which is a

continuation of the labyrinth’s tubuli reaching as far as the supra-esophageal ganglion. These

tubuli eventually return to the CGC, before flowing into a ventral urinary bladder (VB), the

first part of the terminal ductus. From the VB, a small duct leads to the outside world via the

nephropore (NP). Apart from some ramifications of minor importance, the VB continues

caudally via an extension. This extension of the VB, forms a pre-esophageal connection (PEC)

with the contralateral VB. From this PEC, a square duct (SD) gives rise to the median

compartment (MC). The MC is a voluminous structure, rostro dorsally located to the stomach

and can be subdivided into two lobes (a ventral and dorsal lobe). To the lateral side of the

esophagus, the MC continues as the lateral compartment (LC) laterally flanking the gut. The

LC is in its turn flanked by the large protocephalon adductor. The extension of the VB, which

gives rise to the MC, is also connected to the LC at the lateral side of the esophagus. Caudally,

the LC diminishes in height to form the caudal extensions (CE), which end blindly at the height


81

of the hepatopancreas. Along the entire structure of the antennal gland, several organs are

in close proximity of the excretory organ: the heart, hepatopancreas, oesophagus, stomach,

gut, brain, and lymphoid organ. Even more so, many ramifications of the structure were

found, intensifying the contact between the antennal gland and the aforementioned organs,

as well as muscles.


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Fig. 1 The general morphology of the nephrocomplex. A. Lateral view, B. Dorsal view, C. Lateral view in the

median compartment and the ventral bladder, D. schematical overview and orientation of the antennal gland.

RC: rostral compartment; CGC: compact glandular part; VB: ventral bladder; MC: median compartment; DL:

dorsal lobe of the MC; VL: ventral lobe of the MC; LC: lateral compartment; CE: caudal extensions; POC: pre-

esophageal connection; SD: square duct; NP: nephropore; Oe: esophagus; S: stomach; N: nerve; HP:

hepatopancreas

3.4.2. Compact glandular compartment

Upon manual dissection of the antennal peduncle, a clean, white, bean-shaped structure

becomes visible. This structure is bilaterally present and located at the transition from

peduncle to cephalothorax and was named the compact glandular compartment of the

antennal gland (CGC). This connects to a urinary bladder, only visible on a crosscut of the

unopened hemocoel. Depending on the size of the animal, this urinary bladder can be

visualized either by bright-field microscopy or the naked eye.

Central in the CGC, the hemolymph-filtering coelomosac can be found, surrounded by the

filtrate-modifying tubuli known as the labyrinth and by hemolymph spaces (Fig. 2). The

hemolymph sinuses attach to the coelomosac domes (see further) and opposite of the

coelomosac side, they possess a flattened cell layer. The coelomosac itself comprises a central

lumen lined by podocytes with big, irregular nuclei, little cytoplasm, and many vacuoles. The

lumen of the coelomosac consists of big spaces and bow-shaped coelomosac domes. The

antennal artery penetrates the compact glandular compartment and runs in close connection

to the coelomosac and hemolymph spaces. The coelomosac lumens will gather centrally in

the compact glandular compartment and merge to one big vertical coelomosac lumen. This

large central lumen will gradually split into several smaller tubuli to form the labyrinth. The

tubuli of the labyrinth bathe in the hemolymph and their cells are cubical to columnar. They

possess a brush border, and the nuclei are basally located, regular and smaller compared to

the coelomosac podocytes. The cytoplasm is slightly lighter in colour than the cytoplasm of

the coelomosac podocytes. As mentioned before, the tubuli of the labyrinth are located

around the coelomosac. In between the coelomosac and the tubuli of the labyrinth, the

hemolymph sinuses can be found. At the ventral side of the compact glandular compartment,

the labyrinth will eventually debouch into the ventral urinary bladder.


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Fig 2. Overview and details of the compact glandular compartment. A. 3D-reconstruction of the compact

glandular compartment (CGC), B. Cellular morphology of the coelomosac lumen (c*), encompassed by

coelomosac domes (cd) and hemolymph sinuses (hs) (HE-staining), C. Schematic overview of the flow circulation

inside the compact glandular compartment, aA: antennal arteria; cc: central coelomosac lumen; NP:

nephropore; L: Labyrinth; VB: ventral bladder, D. HE-Histological blueprint of the compact glandular

compartment. The coelomosac (c) is surrounded by the labyrinth (L) and together they make up the compact

glandular compartment. The black arrowhead points to the rostrum.

The compact glandular part is penetrated by the antennal artery. The main branch is

surrounded by a dense connective tissue layer, but smaller branches towards the direction of

the coelomosac lose their connective tissue layer. The coelomosac lumen was not observed

to make direct contact to the antennal artery. Rather, the antennal artery ends in hemolymph

sinuses, overlapping the coelomosac domes.

At the medial side of the labyrinth, a small longitudinal muscle can be found. The dorsal origin

of this muscle is at the inside of the exoskeleton and a small part penetrates the compact


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glandular part. The muscle runs medially from and is connected to the labyrinth before ending

ventrally inside the wall of the ventral bladder.

3.4.3. The rostral compartment

The tubuli of the labyrinth inside the compact glandular compartment do not just run to the

ventral bladder but spread out rostrally to form the rostral compartment. At first, they run

along the basipodite and the coxipodite of the first and second antennal pair. Eventually, they

gradually increase in number and start filling the entire cephalon, even surrounding the supra-

esophageal ganglion. Here, tubuli of both sides are seemingly connected with each other,

thus forming one rostral compartment of the antennal gland. When traced back, these tubuli

return to the compact glandular part. At the height of the rostral structure, the tubuli are

composed out of a monolayer of cubical cells with large, round nuclei and have a light pink

cytoplasm on HE-staining. The main branch of the antennal artery, after penetrating and

leaving the compact glandular compartment, continues rostrally but keeps its distances from

the tubuli of the rostral compartment. One exception occurs at the height of the

protocephalon, where one layer of tubuli encloses the antennal artery shortly (Fig.3).


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Fig. 3. Cross section of the cephalon. The rostral compartment fills the hemocoel. A. The tubuli of the rostral

compartment (TRC) are spread throughout the hemocoel of the cephalon. aA: antennal artery; *: detail of the

association between the antennal artery and the rostral compartment; CT: connective tissue; HC: hemocoel; N:

nerve; PA: protocephalon attractor. B. The rostral compartment surrounds the supraesophageal ganglion (SEG).


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3.4.4. The ventral bladder

The tubuli of the rostral compartment eventually return to the compact glandular

compartment (CGC), where they end up in the bilaterally present ventral urinary bladder (VB).

Connections with the CGC can occur at a variable number of places. Contrary to most other

parts of the antennal gland, the bladder wall is made up of multiple cell layers. It is topped by

a single squamous epithelium with bulging nuclei and a microvilli brush border. Underneath,

a lamina muscularis above loose connective tissue can be found. Finally, a lamina serosa

defines the outer surface of the VB. Sometimes, the epithelium can be observed to be two

cell layers thick. To the caudal side, the VB has an extension which has a narrow lumen and

splits into three structures (Fig. 4). Medially, a connection with the contralateral VB is made

in the form of the pre-esophageal connection (PEC). To the caudal side, the extension of the

VB ends in the lateral compartment (LC). On the basal side and around the extension of the

VB, ramifications of unknown significance can be found. Lastly, the VB has a short duct via

which the evacuation of urine takes place. Departing from the VB, this duct starts triangle-

formed, changing into a square, ending as a moon-shaped duct before connecting to the

outside world via the nephropore.

Fig. 4. 3D-reconstruction of the ventral bladder and its continuations. A. lateral view on the ventral bladder

(VB). NP: Nephropore, VBE: ventral bladder extension. B. Rostral view on VB, compact glandular compartment

(CGC), lateral compartment (LC), and median compartment (MC) with ventral lobe (VL) and dorsal lobe (DL).

PEC: pre-esophageal connection; SD: square duct; GMA: gastric mill attractor muscle.


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Around the VB, three important muscles can be found (Fig. 5), i.e., the coxipodite adductor

(CA), the small, and the large scaphocerite abductor (SSA and LSA). Both the CA and the SSA,

originate from the LSA. The LSA muscle attaches in the scaphocerite, to run through the

coxipodite. Just rostrally from the VB, the SSA and the CA branches split from the main muscle.

The SSA runs lateral from the VB and the CA runs medially, while the LSA lays ventrally from

the VB. All three muscles end on the inside of the exoskeleton, caudally from the VB. The

urinary ventral bladder is thus positioned between the coxipodite adductor (CA), the minor

scaphocerite abductor (SSA) and the major scaphocerite abductor (LSA). Furthermore, the

transition between the ventral bladder and the terminal duct is jammed between extensions

of the same two muscles, namely the coxipodite adductor muscle (dorsal) and the large

scaphocerite abductor muscle (lateroventral).

Fig. 5. Muscles associated with the ventral bladder. VB: ventral bladder; LSA: large scaphocerite abductor; SSA:

small scaphocerite abductor; CA: coxipodite adductor; CGC: compact glandular compartment; VBE: ventral

bladder extension; VB*: ramifications of the VB; n: nerve; E: exoskeleton.

3.4.5. The median compartment

As mentioned before, the extension of the ventral bladder has a pre-esophageal connection

(PEC) with the contralateral VB. From the PEC, a square duct runs dorsally giving rise to the

median compartment (Fig 1C and Fig 4B). The MC is a large bladder structure, covering the

stomach rostro dorsally. It can be divided in two parts: a ventral lobe (VL) and a dorsal lobe

(DL). These lobes are parted by the anterior gastric mill attractor muscles. Caudally, the


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structure will gradually turn in the lateral compartment (LC, Fig 1C). As is the case with the

VB, the VB too is associated with several muscles. Beside the aforementioned anterior gastric

mill attractor separating and defining the two lobes, the large protocephalon attractor runs

laterally from the MC and the epistomal stator muscle runs lateral from the DL, which is also

dorsally covered by two posterior protocephalon levator muscles (Fig. 6)

Fig 6. The muscles associated with the median compartment. DL: dorsal lobe of the median compartment; VL:

ventral lobe of the median compartment; PA: protocephalon attractor muscle; ES: epistomal stator muscle;

GMA: gastric mill attractor muscle; S: stomach. The black arrowhead points rostrally.

3.4.6. The lateral compartment with the caudal extensions

Bilaterally from the esophagus, a big longitudinal bladder structure rises: the lateral

compartment (LC, Fig. 1). The lateral compartment is connected with the extension of the

ventral bladder (VBE) and to both the ventral and the dorsal lobe of the median compartment

(MC). The two halves connect via a post-esophageal connection, similar to the pre-esophageal

connection. Laterally flanking the esophagus, stomach and gut, the LC runs caudally, gradually

decreasing in height. In its turn the LC Is flanked by the two large protocephalon attractor

muscles (Fig 7). The epithelium of the LC contains only one cell layer of flattened epithelium.

The LC is attached to both the stomach and the protocephalon attractor muscle. Occasionally,

a small muscle layer can be observed. This structure has only one cell layer of flat epithelium

and is attached to the large protocephalon attractor muscle on the lateral side. Sometimes a


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small muscle layer can be observed between the large protocephalon attractor muscle and

the lateral compartment. The upper part of the lateral compartment lays on top of the

stomach. Here the cells of the antennal gland are not connected to surrounding tissue but lay

loose in the haematocele and consists of an epithelial monolayer and a consistent smooth

muscle layer. To the caudal side, the LC continues as the caudal extensions (CE) dorsally

limited by the pylorus stomach and ventrally by muscle layers and/or exoskeleton. Just

rostrally of the hepatopancreas, a last connection to the contralateral side can be found. Next,

de CE will diverge around the hepatopancreas, ending blindly. These are the most caudal

structures of the antennal gland.

Besides the apparent protocephalon attractor muscles, there are other muscles associated

with the LC (Fig. 7). The esophageal dilators attach to the inside of the exoskeleton and line

the lateral compartment ventrally and dorsally at the height of the esophagus, where these

muscle group ends. Next, the maxillary tensor muscles connect the ventral side of the

stomach to the inside of the exoskeleton and trap parts of the LC between those structures.

Similarly, two previously undescribed oblique muscles attach to both the stomach and the

exoskeleton. However, these two muscles are much thinner and are located medially from

the mandibulary tensor muscles. Unlike the latter, these muscle strains do not have a straight

path, but a rather oblique one. Finally, the mandibular adductor muscles have the same

conformation but are more caudally located.


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Fig. 7. Muscles associated with the lateral compartment (LC) and caudal extensions (CE). A. Frontal view on a

cross section of the LC, B. Ventral view. PA. protocephalon attractor; ED. esophageal dilator; S. stomach; n.

nerve. B. Ventral view on LC and EC. PoEC. post-esophageal connection; MT. maxillary tensor; MA. mandibular

adductor; OM. oblique muscle; HP. hepatopancreas.

3.5. DISCUSSION

3.5.1. Anatomy of the antennal gland based on HE-staining

In this study, the anatomy of the antennal gland was explored in full. Besides the already

known structures (Bell and Lightner, 1988), the nephropore, terminal duct with urinary

bladder, labyrinth, and coelomosac (forming the compact glandular compartment, CGC), we

have additionally uncovered a rostral continuation of the labyrinth surrounding the supra-

oesophageal ganglion, coined as the rostral compartment (RC). The ventral urinary bladder

(VB) appears to possess several continuations: (i) a terminal duct leading to the nephropore

(NP) and ultimately to the outside world, (ii) the pre-esophageal connection (PEC) connecting

the contralateral urinary bladder and (iii) the lateral compartments (LC), two bilateral large

sheets flanking the stomach. These lateral compartments flow into the caudal extensions (CE),

ending blindly at the height of the hepatopancreas. Rostrally from the stomach, the lateral

compartments are connected with each other, forming a median compartment (MC). The two

large protocephalon attractor muscles subdivide the median compartment into a ventral and

a dorsal lobe.

In 1959, Young already described colorations above and below the supra-esophageal ganglion

after introducing dye in the nephropore using Penaeus setiferus as subject. We are convinced


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these colourations are the rostral compartment. We also hypothesize that the rostral

compartment is homologous to the loops of Henle as found in mammalian kidneys. Nakamura

and Nishigaki described a larger and more complex anatomy than predecessors, using a clay

model in Penaeus japonicus. However, they stated that there is only one continuation of the

ventral urinary bladder, the coelomosac (Nakamura and Nishigaki, 1991; Nakamura and

Nakashima, 1992). Based on the histology, we disproved this theory. We found that the

coelomosac is surrounded by the labyrinth inside the CGC and that the VB is a continuation

of this labyrinth in accordance with other descriptions of the penaeid antennal gland (Xiaoyun

et al., 2003). Furthermore, many of the structures described in their paper are different or

have an altered configuration in comparison to this study performed in P. vannamei. They

also described ramifications coming from the VB. Here, this conformation is confirmed. In

addition, these ramifications could be found all over the antennal gland.

A voluminous median compartment with a division in a ventral and dorsal lobe has not yet

been described. Nakamura and Nakashima (1992) did come close with their clay model

reconstruction of P. japonicus larvae’s antennal gland. They presented a “median lobe of the

transversal coelomosac” at the site where this chapter describes the median compartment.

The denomination of coelomosac is very likely incorrect as the histology of the median

compartment and the actual coelomosac inside the compact glandular part do not match.

Here, the cells do not have a cell morphology that can be linked to filtration or filtrate

modifying functions. They are rather adapted to a more dynamic function. The same goes for

other parts of their clay model. The overall structure of the median compartment described

in this chapter and the “median lobe of the transversal coelomosac”, which they describe as

a “thin longitudinal dorsal structure” diverse significantly. Quite possibly, this difference can

be explained by the use of animals. The Nakamura study did not only use slightly different

species (P. japonicus vs P. vannamei), but they also used 0.7g larvae, while this chapter used

post-larvae of ca. 9g indicating that the structure reported by the first could be an immature

and undeveloped version of the median compartment in post-larvae. However, this raises the

question on how the newly described anatomy evolves during subsequent life stages. More

research hereon could be beneficial.


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The quality of the histology-based 3D-model may seem under par but this is due to the

presence of a thick cuticular shield (the carapax) which makes slicing ultrathin cross-sections

(<10 µm) extremely difficult. We opted to not remove this carapax because it may have

altered the localization and orientation of some extremely thin antennal gland structures of

interest. The authors noticed that on subsequent slices, the contents were not always

perfectly aligned, leading to a lower 3D quality. We theorize that the pressure of the knife on

the tough carapax may have induced minor changes to the underlaying tissues, slightly

mismatching the alignment of serial sections.

Finally, the unravelling of the complete structure indicates that the antennal gland is a perfect

portal for pathogen entry. Its wide distribution in the shrimp cephalothorax results in close

contact to all WSSV-susceptible organs: the nervous system, the alimentary tract, the

lymphoid organ, the gills, the hepatopancreas, and various muscles. Only the heart is not in

close contact with the antennal gland. Most of the antennal gland’s cell layers are singular

and have no cuticular lining, which allows for a potential rapid entry of pathogens into the

hemolymph.

3.5.2. The flow of filtrate to and through the antennal gland

A plethora of mechanisms of how the coelomosac receives fluid for filtration has been

gathered in the body of literature. This chapter provides novel insights into this matter.

According to the studies of Xiaoyun et al. (2003) and Buranajitpirom et al. (2010), this chapter

situates the coelomosac in the middle of the antennal gland’s filtration structure (here named

the compact glandular compartment) surrounded by the tubuli of the labyrinth. Based on the

histology, we propose the following model for hemolymph filtration: the antennal artery

penetrates the compact glandular compartment. Inside the CGC, the antennal artery is still

surrounded by layers of connective tissue. Further inside the CGC, at the height of the

coelomosac, this connective tissue layer disappears and ramifications of this artery end in

hemolymph sinuses. These sinuses surround coelomosac domes. This description of the

filtration flow is complementary to findings in several other crustacean antennal or green

glands e.g. palaemonids (Panikkar, 1941), fiddler crab (Schmidt-Nielsen et al., 1968),

Procambarus (Schaffner and Rodewald, 1978), Crangon crangon (Tikær Andersen and


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Baatrup, 1988), Macrobrachium rosenbergii (Al-Mohsen, 2009), Astacus (Khodabandeh et al.,

2005), and the semi-terrestrial crab ocypode (Tsai and Lin, 2014), but is in contrast to the

studies of Holliday and Miller, 1984 on decapod Crustaceans and Xiaoyun et al., 2003 on

Penaeid shrimp where a direct contact of the coelomosac to the antennal arterial wall is

described. Next, we described the coelomosac dome’s lumen to be in direct contact to a larger

central coelomosac lumen which in turn, debouches into one large collection tube. It is from

here, that the tubuli of the labyrinth are formed.

The tubuli of the labyrinth thus surround the coelomosac and make up the body of the

compact glandular compartment. We did not observe different labyrinth epithelium to divide

it into a labyrinth I and II like Al-Mohsen (2009) and Khodabandeh et al. (2005). The long

columnar epithelium with microvilli brush borders and large basal nuclei resembles active

cells involved in modifying the primo filtrate. This is similar to the descriptions of Tsai and Lin

(2014) and khodabandeh (2005).

This study describes a large body of tubuli filling the hemocoel of the cephalon and

surrounding the supraesophageal ganglion (brain). This structure was named the ‘rostral

compartment’. The tubuli of the rostral compartment possess a very similar epithelium as the

labyrinth. Moreover, the lumina of the tubuli of the labyrinth are in direct contact to the tubuli

of the rostral compartment. Functionally, it is plausible that the rostral compartment is part

of the labyrinth or at least an extension or continuation thereof. The tubuli of the labyrinth

inside the compact glandular compartment would spread rostrally (to the rostral

compartment) and occupy a large part of the cephalon. This conformation could benefit from

the resulting surface enlargement to provide more filtrate modifying opportunities. It is

described that the antennal gland of Penaeids regulates the hemolymph volume and its ion

balance (Lin et al., 2000). The large surface of the rostral compartment would aid the shrimp

in quickly regulating hemolymph volume and/ or pressure during periods of osmostress.

Further functional research on these tubuli is warranted.

Finally, the tubuli of the labyrinth flow into the ventral urinary bladder. From here, the

contents can be evacuated to the outer world via the nephropore (mictio) or can be passed


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on to the lateral compartments and or median compartment. Muscles surrounding these

structures could regulate and determine the direction of the content flow (see further).

In summary, the hemolymph is delivered to the coelomosac by the antennal artery ending in

several hemolymph sinuses. These sinuses are situated around the coelomosac. It is here that

the hemolymph is filtrated, i.e., during the transition from hemolymph sinus to coelomosac

dome. These domes are in direct contact to a few central coelomosac lumens, which end in

one big collection space. From here, the tubuli of the labyrinth sprout. Much like the loop of

Henle in mammals, these tubuli branch out to the cephalon (the rostral compartment) and

return to the labyrinth, finally debouching into the ventral bladder whereafter the contents

can be passed to other parts of the antennal gland or expulsed outside the animal via the

nephropore.

3.5.3. Muscles associated with the antennal gland and their functions

The VB is surrounded by three major muscles (Fig 5): medially the coxipodite adductor and

the small scaphocerite abductor (SSA), ventrally the large scaphocerite abductor (LSA). The

scaphocerite adductor muscles indubitably play a major role in squeezing the VB to empty its

contents, aided by the smooth muscle layers residing in the bladder wall. The transition

between the VB and this terminal duct is jammed between the coxipodite adductor (dorsal)

and the large scaphocerite abductor (lateroventral). No other muscular structure was found

capable of performing a sphincter-like function. Therefore, we assume that the urinary

bladder is sealed by a combination of the two latter muscles and the nephropore. This links

scaphocerite abduction to the mictio process.

The new structures of the antennal gland whereby compartments were retrieved caudally

from the ventral urinary bladder, raises the suspicion that an anterograde flow may be

possible. When the abductors of the scaphocerite (SSA and LSA) contract together with the

adductor of the antennal coxipodite (CA), the terminal ductus will be pinched between these

muscles and a urinary flow towards the median compartment and the lateral compartment

could occur.


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The MC is divided into a VL and DL by the anterior gastric mill attractor muscle. The large

protocephalon attractor runs laterally from the MC and the epistomal stator muscle runs

lateral from the DL, which is also dorsally covered by two posterior protocephalon levator

muscles. Again, a coordinated action from these muscles could modulate the contents and

shape of the MC. Histologically, the MC is also similar to the VB, but with a slightly thicker

smooth muscle layer, hinting at a dynamic function.

The large protocephalon attractor muscle is not only associated with the median

compartment, but also with the lateral compartment. Other muscles associated with the LC

are the esophageal dilators, the maxillary tensor muscles, mandibular adductor and two

oblique muscles. All these muscles have the potential to isolate parts of the LC, meaning that

the shrimp can potentially regulate the volume of the LC and its caudal extensions. Two of

these muscles, the two oblique muscles have never been described before. There

conformation resembles that of the cruciate ligaments in mammalian knee articulations,

squeezing the caudal extensions between the inside of the exoskeleton and the outside of the

stomach/gut. When all these muscles coordinate, the lateral compartment can be

compartmentalised and therefore urine could be pumped towards the caudal extensions and,

if necessary, back to the ventral bladder in a controlled fashion.

3.5.4. Possible mechanical role of the antennal gland in moulting

The position and possible functions of the abovementioned muscles, together with the

localisation of the median compartment and the caudal extensions feeds suspicion that the

antennal gland could hypothetically play a pivotal role in the mechanical process of moulting.

When the discussed muscles cooperate, the median compartment and/or caudal extensions

of the lateral compartment could be actively filled with fluids, either through increased

filtration of hemolymph (and peroral uptake of water) or (less likely) directly from ambient

water. Filled median and lateral compartments could result in an increase of pressure on the

inside of the carapax roof. When one observes the ecdysis of decapods, it becomes apparent

that the transition between the cephalothorax (covered by the carapax) and the abdomen is

the place where the animal brakes out of its old shell. In shrimp, the cephalothorax (and thus

also the carapax) is connected to the abdominal exoskeleton via a fragile hymen. Pressure on

this hymen is necessary to make it rupture and thereby separating carapax from abdomen.


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Modulation of the volume of the caudal extensions and/or median compartment could

increase the internal pressure on the inside of the carapax roof enough to break the hymen

when the shrimp violently contracts the body during ecdysis (the act of moulting).

3.6. CONCLUSION

In conclusion, the antennal gland has a substantially wider distribution throughout the

cephalothorax and is a much more complex structure than previously assumed. Because of

the vast distribution of the organ in the cephalothorax, not limited to the antennal peduncle,

the antennal gland’s function as an excretory organ, and the complex of diverticles in

connection with the bladders, we propose a new name for this organ: the nephrocomplex.

The prefix ‘nephro’ from the Greek ‘nephros’ for kidney and the suffix ‘complex’ because of

the large numbers of diverse subunits that the excretory organ consists of. Furthermore, its

anatomy, morphology, and cellular structure are optimal for the excretion organ to be a

pathogen entry portal: (i) there is a complete absence of cuticular or chitinous lining along

the lumen of the organ, (ii) the thickness of the majority of the organ is limited to one or two

cell layers, allowing for a potentially fast spreading of WSSV to the hemolymph and (iii) its

association to many WSSV-susceptible organs by proximity and/or the presence of many

ramifications of the nephrocomplex. Additionally, this chapter indicates that the

nephrocomplex could be involved in the moulting process. Since links between moulting and

WSSV-infection where laid, it would be profitable to investigate this matter in depth.


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3.7. REFERENCES

Al-Mohsen, I., 2009. Macrobrachium rosenbergii (de Man 1879), The antennal gland and the role of pheromones in mating behaviour. Doctoraatsthesis, Institute of Aquaculture, University of stirling, 60-120, Institute of Aquaculture. University of Stirling, Stirling, Scotland. Anderson, D. T., 1973, Crustaceans, in D. T. Anderson (ed.), Embryology and Phylogeny in Annelids and Arthropods (Pergamon), 263-364. Bell, T.A. and Lightner, D. A., 1988, A handbook of normal penaeid shrimp histology (Baton Rouge, LA, USA: World Aquaculture Society). Buranajitpirom, D., et al., 2010, Adaptation of the black tiger shrimp, Penaeus monodon, to different salinities through an excretory function of the antennal gland, Cell and Tissue Research 340 (3), 481-89. Chang, P. S., et al., 1996, Identification of white spot syndrome associated baculovirus (WSBV) target organs in the shrimp Penaeus monodon by in situ hybridization, Diseases of Aquatic Organisms, 27 (2), 131-39. Corteel, Mathias, et al., 2009, Molt stage and cuticle damage influence white spot syndrome virus immersion infection in penaeid shrimp, Veterinary Microbiology, 137 (3-4), 209-16. Corteel, M., et al., 2012, Moult cycle of laboratory raised Penaeus (Litopenaeus) vannamei and P. monodon, Aquaculture International 20 (1), 13-18. Domínguez-Borbor, C., et al., 2019, An effective white spot syndrome virus challenge test for cultured shrimp using different biomass of the infected papilla, MethodsX, 6, 1617-26. Escobedo-Bonilla, C. M., et al., 2008, A review on the morphology, molecular characterization, morphogenesis and pathogenesis of white spot syndrome virus, Journal of Fish Diseases, 31 (1), 1-18. Felgenhaur, BE, 1992, Decapod Internal Anatomy, in W. Harrison Frederick and E. Ruppert Edward (eds), Microscopic anatomy of invertebrates: Decapod Crustacea 10, 7-43, Wiley-Lis, New York. Freire, Carolina A., Onken, Horst, and McNamara, John C., 2008, A structure–function analysis of ion transport in crustacean gills and excretory organs, Comparative Biochemistry and Physiology Part A: Molecular & Integrative Physiology, 151 (3), 272-304. Holliday, Charles W. and Miller, David S., 1984, Cellular Mechanisms of Organic Anion Transport in Crustacean Renal Tissue, American Zoologist, 24 (1), 275-84.


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Icely, J. D. and Nott, J. A., 1979, The general morphology and fine structure of the antennary gland of Corophium volutator (Amphipoda: Crustacea), Journal of the Marine Biological Association of the United Kingdom, 59 (03), 745-55. Khodabandeh, S., Charmantier, G., and Charmantier-Daures, M., 2005, Ultrastructural studies and Na+, K+-ATPase immunolocalization in the antennal urinary glands of the lobster Homarus gammarus (Crustacea, Decapoda), Journal of Histochemistry and Cytochemistry, 53 (10), 1203-14. Lin, Shian-Chuann, Liou, Chyng-Hwa, and Cheng, Jin-Hua, 2000, The role of the antennal glands in ion and body volume regulation of cannulated Penaeus monodon reared in various salinity conditions, Comparative Biochemistry and Physiology Part A: Molecular & Integrative Physiology, 127 (2), 121-29. Nakamura, K. and Nishigaki, Y., 1991, Structure of Antennal Gland in Kuruma Prawn Penaeus-Japonicus, Nippon Suisan Gakkaishi, 57 (10), 1859-63. Nakamura, K. and Nakashima, N., 1992, Structure of Antennal Gland Coelomasac in the Kuruma Prawn, Nippon Suisan Gakkaishi, 58 (8), 1551-51. Panikkar, N.K., 1941, Osmoregulation in some palaemonid prawns. Journal of Marine Biology Association 25, 317 – 359. Pina, S., Russell-Pinto, F., Rodrigues, P., 2007, Clarification of Cercaria sevillana (Digenea: Microcephallidae) life cycle using morphological and molecular data. Journal of Parasitology 93, 318–322. Peterson, Darryl R. and Loizzi, Robert F., 1974, Ultrastructure of the crayfish kidney—coelomosac, labyrinth, nephridial canal, Journal of Morphology, 142 (3), 241-63. Riegel, J. A., 1966a, Analysis of Formed Bodies in Urine Removed from Crayfish Antennal Gland by Micropuncture, Journal of Experimental Biology, 44 (2), 387-&. Riegel, J.A., 1966b, Micropuncture Studies of Formed-Body Secretion by Excretory Organs of Crayfish Frog and Stick Insect, Journal of Experimental Biology, 44 (2), 379-&. Riegel, J.A., 1970, A New Model of Transepithelial Fluid Movement with Detailed Application to Fluid Movement in Crayfish Antennal Gland, Comparative Biochemistry and Physiology, 36 (2), 403-&. Rodríguez, J., Echeverría, F., Maldonado, M., Blake, S., Balladares, A., Ruiz, J., 2009. Data from El camarón Penaeus vannamei y sus patógenos virales, estado actual de conocimientos de mecanismos de defensa celular. ESPOL Repository. Internet reference: http://www.dspace.espol.edu.ec/xmlui/handle/123456789/4723 (consulted on 21 March 2021), Deposited 11 March 2009.


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Ryazanova, T.V., Eliseikina, M.G., Kalabekov, I.M., Odintsova, N.A., 2015. A herpes-like virus in king crabs: Characterization and transmission under laboratory conditions. Journal of Invertebrate Pathology 127, 21-31. Schaffner, A., Rodewald, R., 1978, Filtration barriers in the coelomic sac of the crayfish, Procambarus clarkii. Journal of ultrastructure research 65, 36-47. Schmidt-Nielsen, B., Gertz, K.H., Davis, L.E., 1968, Excretion and ultrastructure of the antennal gland of the fiddler crab Uca mordax. Journal of Morphology 125, 473-496. Thrupp, t.J., Lynch, S.A., C., W.E., Malham, S.K., Vogan, C.L., Culloty, S.C., Rowley, A.F., 2013, Infection of juvenile edible crabs, Cancer pagurus by a haplosporidian-like parasite. Journal of Invertabrate Pathology 114, 92-99. Thuong, K.V., Tuan, V.V., Li, W.F., Sorgeloos, P., Bossier, P., Nauwynck, H., 2016, Per os infectivity of white spot syndrome virus (WSSV) in white-legged shrimp (Litopenaeus vannamei) and role of peritrophic membrane, Veterinary Research, 47, 12. Thuong, K.V., Tuan, V.V., Li, W., Sorgeloos, P., Bossier, P., Nauwynck, H., 2016, Effects of acute change in salinity and moulting on the infection of white leg shrimp (Penaeus vannamei) with white spot syndrome virus upon immersion challenge. Journal of Fish Diseases 39, 1403-1412. Tikær Andersen, J., Baatrup, E., 1988, Ultrastructural localization of mercury accumulations in the gills, hepatopancreas, midgut, and antennal glands of the brown shrimp, Crangon. Aquatic Toxicology 13, 309-324. Tsai, J.R., Lin, H.C., 2014, Functional anatomy and ion regulatory mechanisms of the antennal gland in a semi-terrestrial crab, Ocypode stimpsoni. Biol Open 3, 409-417. Vogt, G. (2002), Functional Anatomy, in D.M. Holdich (ed), Biology of Freshwater Crayfish, 53-151, Blackwell Science ltd, Oxford, UK. Wang, Y. C., et al., 1998, Experimental infection of white spot baculovirus in some cultured and wild decapods in Taiwan, Aquaculture, 164 (1-4), 221-31. Xiaoyun, L., Wei, X., and Zhenmin, B., 2003, Histology and Functions Study of the Antennal Gland of Penaeus chinensis, Journal of Ocean University of Qingdao, 33 (6), 854 – 60. Xugang, He, Guangfu, Hu, and Guangtao, Lu, 2013, The effect of lower salinity on microstructure of antennary gland of Litopenaeus vannamei, Journal of Food, Agriculture & Environment, 11 (1), 782-85. Young, Joseph, 1959, Morphology of The White Shrimp Penaeus setiferus (Linnaeus 175), Fishery Bulletin 145 (59), 173.

Chapter 4

µMRI imaging of Penaeus vannamei: the involvement of the shrimp nephrocomplex during the moulting

process

G.M.A. De Gryse





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4.1. ABSTRACT

For several decades, the shrimp industry has been severely terrorised by the white spot

syndrome virus (WSSV). The development of decent preventive and prophylactic measures

have been hampered by a lack of pathogenesis knowledge. Especially the place of WSSV-entry

has been the subject of heavy debate in recent years. Based on the anatomy and morphology

of the nephrocomplex, this organ is now in pole position to be named the major entry portal

for WSSV. Not only does the structure lacks an inner cuticular lining, but it is also vastly spread

throughout the cephalothorax and its cell layer is only a few cells thick. Also, osmostress, a

primary function of the nephrocomplex, has been linked to WSSV-infection. In the previous

chapter, a link to the moulting cycle was made. The moulting cycle is also linked to WSSV-

infection. Here, micro-Magnetic Resonance Imaging (µMRI) was applied to live Penaeus

vannamei to both confirm the histology-based 3D-reconstruction made in chapter 3, and to

further investigate the role of the nephrocomplex during the moulting process of penaeid

shrimp. Due to the high intensity returned by the high-water content in urine, clear images of

the nephrocomplex could be made with µMRI. Interesting conclusions concerning the

moulting and growing process resulted from this methodology. Furthermore, applying µMRI

to live shrimp could be a powerful tool to examine the in vivo anatomy of the internal organs.

Key words: Penaeus vannamei, antennal gland, nephrocomplex, µMRI, moulting

Chapter 4 µMRI

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4.2. INTRODUCTION

Since the discovery of the WSSV, the debate on the entry portal of the virus is ongoing.

Although the scientific consensus is fixed on the peroral route (Corteel et al., 2009), the

efficiency of the infection route is very low and difficult to reproduce (Perez et al., 2005;

Laramore, 2007). The presence of a protective chitinous layer covering the inside of the

alimentary tract which protects the animal against viral invasion and the fact that the tubular

epithelial cells are refractory to a WSSV infection, increases the possibility for another major

entry portal (Young, 1959; Hegedus et al., 2009; Thuong et al., 2016). Therefore, the search

for the entry portal was redirected to structures not protected by such an impermeable layer.

In Chapter 3, it was evidenced ex vivo that the nephrocomplex (formerly known as the

antennal gland) of Penaeus vannamei not only lacks a cuticular lining, but also has an

extremely complicated anatomy. Besides the long known coelomosac, labyrinth, bladder and

exit duct, a series of bladder compartments were retrieved caudally of the aforementioned

structures. In vivo confirmation of the 3D-model based on HE-sections would anchor the

newly found morphology of the organ.

Moreover, the localisation of these compartments (especially the dorsal lobe of the median

compartment and the caudal extensions of the lateral compartment) in combination with

several associated muscles, suggested that the volume of these bladders could be actively

modulated and possibly recruited during the moulting process. The nephrocomplex is already

known to be involved in the moulting process through modulation of several moulting

hormones (Chang, 1985; Corteel and Nauwynck, 2010; Mykles, 2011), the metabolism of

calcium needed for the mineralisation of the new exoskeleton (Ahearn et al., 2004; Wheatly

et al., 2004), and possibly the volume regulation of the shrimp’s body (Lin et al., 2000).

Previously, links have been established between moulting and increased susceptibility of

shrimp towards viral infections (Lightner et al., 1983; Corteel et al., 2009; Van Thuong et al.,

2016). Confirmation of an active role of the shrimp’s excretion organ during the moulting

process would thus further strengthen the evidence towards the nephrocomplex as a major

viral entry portal.

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4.3. MATERIAL AND METHODS

4.3.1. IN VIVO 3D-RECONSTRUCTION OF THE ANTENNAL GLAND USING µMRI

For the in vivo confirmation of the antennal gland 3D-reconstruction based on HE-stained

coupes, we applied the µMRI-technology of the Infinity lab (Faculty of Medicine and Health

Sciences, Ghent University, Belgium), generally employed for laboratory rats in the study of

brain tumours.

Animals

Shrimp originating from Hawaii were collected at the Artemia Reference Center (ARC) and

transported to the Laboratory of Virology, Faculty of Veterinary Medicine, Ghent University

for acclimatisation. There, 25 shrimp were screened at different moulting stages (A, B, C, D1

and D2) according to the method of Corteel et al. (2012) and individually transported to the

Infinity lab of the UZ Ghent, where µMRI was performed.

Image acquisition

Magnetic resonance images were acquired on a 7 Tesla small animal MRI (Bruker BioSpin

PharmaScan® 70/16, Ettlingen, Germany) using a transmit/receive volume coil with 40 mm

inner diameter. A T2-weighted TurboRARE sequence was acquired using the following

settings: repetition time (TR) 6105 ms, echo time (TE) 37 ms, in-plane resolution 109 µm, 50

contiguous transverse slices of 600 µm, and a field-of-view of 30x25 mm, resulting in a total

acquisition time of 17 minutes. Before commencing the MRI procedure, the live shrimp were

introduced into a modified falcon tube (50 mL) for fixation. The falcon tube was surrounded

by a spiral of PVC tubing. To keep the shrimp immobile and alive during the entire duration

of the image acquisition, the ambient temperature inside the falcon tube was reduced to 4°C,

by running ice-cold water through the tubing using a small electrical pump immersed in ice-

water. Further immobilization was performed using small pieces of flexible foam. Magnetic

resonance images were acquired using a T2-weighted sequence.

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3D-reconstruction & analysis

Using InVesalius software (open-source software, Centro de Technologia da Informação

Renato Archer, Brazil), the structures of interest (antennal gland, alimentary tract and

hepatopancreas) were labelled on sequential 2D-images using threshold values. By stacking

these labels along the Z-axis, a 3D-image could be reconstructed.

To calculate the relative volume of caudal antennal gland parts during different moulting

stages, the area of the antennal gland at the height of the hepatopancreas was calculated

following manual labelling and put into ratio with the area of the hepatopancreas as a

reference. The results were statistically analysed using R (ANOVA and Pairwise t-test with

Bonferroni correction).

4.4. RESULTS

4.4.1. Immobilisation of shrimp by lowering the ambient temperature

Shrimp placed inside the PVC tube while spiralling ice-cold water through the tubing resulted

in docile and immobile shrimp and allowed the shrimp to survive the hour-long µMRI

procedure. The lowering of the temperature also countered the potential harmful heat

generated by the µMRI machine during image acquisition. Hence, the µMRI procedure could

capture the morphology of the nephrocomplex in living shrimp (in vivo).

4.4.2. Differentiation between nephrocomplex and surrounding tissues

Several parts of the nephrocomplex were distinguishable by µMRI. The ventral urinary

bladder, pre-esophageal connection, median compartment, lateral compartment, and the

caudal extensions, were clearly distinguishable on the µMRI images. Because of the watery

content of these structures, a stronger signal on the µMRI was generated by the magnetic

resonance as compared to the surrounding hemolymph and muscle tissue with lower water

content. This resulted in a sharp contrast between the nephrocomplex bladders and the

surrounding tissue. However, the compact glandular compartment and the rostral

Chapter 4 µMRI

107

compartment were only visible on one occasion and with significantly lower contrast. On only

one occasion was the labyrinth and the collection duct of the compact glandular

compartment visible (Fig. 1).

Fig. 1. Single µMRI-image of the shrimp hemocoel. The shrimp compact glandular compartment is visible, along

with the ventral bladder and the ventral lobe of the median compartment. The tubing used to lower the ambient

temperature of the shrimp is also visible on this frame. VB: ventral bladder; CGC: compact glandular

compartment; VL: ventral lobe of the median compartment; M: muscle; HC: hemocoel; and *: the fluid from the

immobilisation device.

4.4.3. In vivo confirmation of the nephrocomplex morphology

Depending on the filling of the structures, the following morphology was observed. The most

rostral structure retrieved on the µMRI was the ventral urinary bladder, including the terminal

duct leading to the nephropore and the connection to the compact glandular compartment.

A connection to the caudal side was found, where it connected to a pre-oesophageal

connection. Caudally from the oesophagus, a post-oesophageal connection was observed.

Chapter 4 µMRI

108

From this connection, one medially positioned compartment, rostro dorsally from the

stomach was identified as the median compartment with a ventral and dorsal lobe including

the hole through which the epistomal stator muscle runs. Another continuation of the

oesophageal connections are the lateral compartments, running alongside the alimentary

tract, to end blindly in a various number of caudal extensions. These caudal extensions could

be retrieved dorsal, ventral and/or lateral to the hepatopancreas (Fig. 2). Other organs were

also visible on the µMRI, such as the brain, muscles, intestinal tract (oesophagus, stomach,

gut), hepatopancreas (with internal structure), and finally, the heart.

Chapter 4 µMRI

109

Fig. 2. 3D-reconstruction of the nephrocomplex based on cross-sections with µMRI. Green: vesicular

compartments of the nephrocomplex; brown: intestinal tract; yellow: hepatopancreas. a-f: µMRI-images

representing cross-sections of the 3D-model. Arrows indicate parts of the nephrocomplex.

Chapter 4 µMRI

110

4.4.4. Volume shifts during different moulting stages

The filling of the caudal extensions during different moulting stages was also investigated with

the µMRI (Fig. 3). The content’s volume of these caudal extensions was compared to the

volume of a ‘household organ’ in analogy of household genes in relative qPCR. The volume of

the hepatopancreas (also visible on the µMRI) was chosen as a household reference because

the size of this organ does not vary over different moulting stages. The contents of the

nephrocomplex’ caudal extensions gradually decreased according to the progressing

moulting stage to reach a minimum in D1 (early pre-moult). In D2 (late pre-moult), the

contents increased substantially. Around ecdysis (stages D2 and A), the volume of the caudal

part was significantly greater compared to the volume observed in animals that were in

intermoult (stages B, C, and D1). The contents of the nephrocomplex in perimoult shrimp

were on average 3.25 times greater when compared to intermoult animals.

3a 3b

Fig. 3. Boxplot with ratio volume of caudal extensions of the antennal gland over volume of the

hepatopancreas measured with µMRI. 3a shows the average volume 3.25 times higher in perimoult than

intermoult. 3b shows the gradual reduction of the volume ratio over the different moulting stages. Stadium: A

to E represent moulting stages A, B, C, D1, and D2.

Chapter 4 µMRI

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4.5. DISCUSSION

4.5.1. µMRI

Never before was the internal anatomy of shrimp studied using (µ)MRI. While the µMRI

procedure was running, shrimp remained calm, and immobile during the whole experiment,

thanks to the in-house immobilisation device which lowered the ambient temperature of the

shrimp. After the experiment, shrimp were still alive, however no test subjects survived longer

than one hour post experiment. Possibly due to a lack of oxygen or increase in stress, or

maybe because the temperature was raised too quickly after the imaging process. The

improvised contraption using spiral PVC-tubing with ice cold water flowing through, managed

to decrease the core temperature of the shrimp, probably inducing some kind of hibernation,

allowing the shrimp to remain quiet and survive the procedure.

The µMRI experiment proved to be successful as the signal from the water content of the

larger parts of the organ was distinguishable from that of the hemolymph caused by the

differences in water content. Overall, the antennal gland followed the same blueprint as

revealed by the histology-based AMIRA reconstruction described in chapter 3, thus,

confirming the histology-based 3D-reconstruction in vivo. Only the rostral compartment and

the compact glandular compartment were not visible on µMRI. On one occasion however, the

µMRI did show a cross section of the compact glandular compartment, but the contrast was

not strong and was never repeated. In case of the compact glandular compartment, we

hypothesized that the contents of these structures resemble the hemolymph too much and

that the volume is too low to be detected using the µMRI-technique. The individual tubuli of

the rostral compartment, which we believe to be the rostral continuation of the glandular

part analogue to the loop of Henle, holds a low volume as well. Combined with the rather

limited resolution of the technique (originally intended for studying rat brains, 10 times the

size of the nephrocomplex) this could explain the non-visibility on the µMRI. The µMRI

reconstruction also revealed the caudal extensions running dorsally and ventrally of the

hepatopancreas in contrast to the histology-based model, where the caudal extensions run

laterally from the hepatopancreas.

Chapter 4 µMRI

112

4.5.2. Moulting

Moulting is a very impactful event on shrimp; drastic changes occur in the shrimp’s body when

the metabolism changes from homeostasis to allostasis while the animal prepares to shed its

old cuticle and grows new tissue. With a complexity and an almost unrivalled extensiveness,

it is not surprising that the nephrocomplex plays a significant role in this process. A first part

is played by metabolising the moulting hormone ecdysone from the Y-organ and excretion of

its metabolites. This way, the organ can regulate the balance between the moult inhibiting

hormone and moulting hormone ecdysone, which determines the progression through the

subsequent moulting stages (Chang, 1985; Mykles, 2011). The moulting hormone ecdysone

reaches a peak in stage D1 (early pre-moult), urging the cells to prepare for ecdysis. Late pre-

moult, these hormone levels drop significantly (Corteel and Nauwynck, 2010). Immediately

before ecdysis, coordinated actions of muscles associated with the nephrocomplex, extra

fluids are pumped to the caudal extensions (CE) of the lateral compartments. The contents of

the CE are now at a maximum. This higher volume aids in raising the inside pressure on the

carapax, putting ample pressure on the ligament between the carapax and the first abdominal

segment. When the shrimp contracts its other muscles, the added pressure by the CE helps

in the shedding of the exuvium. Immediately after the shedding of the old cuticula, the shell

is still soft. This is the only time during the moulting-cycle, when absolute volume of the

shrimp can be increased. Shortly after the ecdysis, the volume inside the CE remains high,

helping shrimp to increase that volume as much as possible while the exoskeleton hardens.

The hardening of the new, still soft shell happens through the process of mineralisation.

Calcium plays a pivotal role in the mineralisation of the new exoskeleton by forming calcite

(CaCO3) and also as an intracellular messenger in the hormonal actions (Ahearn et al., 2004

and Wheatly et al., 2004). Without doubt, the nephrocomplex fulfils an important part in

regulating the calcium concentrations inside the hemolymph, most probably by the cells of

the labyrinth and by the coelomosac and quite possibly by other parts of the organ as well

(e.g., rostral compartment). However, this remains to be investigated. Next, when the shrimp

progress through the moulting stages (A and B), the volume inside the CE will gradually drop,

making room for newly grown tissue. During intermoult (C) and early pre-moult (D1), the

volume inside the CE will be at its lowest and the moulting inhibiting hormone will be in

balance with the moulting hormone ecdysone (Corteel and Nauwynck, 2010). To investigate

the evolution of the CE volume over the different moulting stages, the ratio of the total area

Chapter 4 µMRI

113

of the CE were taken over de total area of the hepatopancreas. This methodology is based on

the assumption that the size of the hepatopancreas stays constant over the different moulting

stages. However, one could argue that the hepatopancreas is a digestive organ and therefore

during periods of starvation could decrease in size. Post moulting is such a period of self-

starvation. However, given that the ratio CE/HP is 3.25 times higher when compared to inter-

moult, such a small potential decrease in size should not have a significant impact on these

results.

4.6. CONCLUSION

The Anatomy and morphology of the shrimp nephrocomplex was confirmed by µMRI in vivo.

Furthermore, this technique led to the discovery of a dynamic function of the nephrocomplex

during the moulting and growing process.

Chapter 4 µMRI

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4.7. REFERENCES

Ahearn, G.A., Mandal, P.K., Mandal, A. (2004). Calcium regulation in crustaceans during the molt cycle: a review and update. Comperative Biochemestry, Part A: Molecular & Integrative Physiology 137, 247-257. Chang, E.S. (1985). Hormonal Control of Molting in Decapod Crustacea. American Zoologist 25, 179-185. Corteel, M., Dantas-Lima, J.J., Wille, M., Alday-Sanz, V., Pensaert, M.B., Sorgeloos, P., Nauwynck, H. (2009). Molt stage and cuticle damage influence white spot syndrome virus immersion infection in penaeid shrimp. Veterinary Microbiology 137, 209-216. Corteel, M., Nauwynck, H.J. (2010). The integument of shrimp, in: Alday-Sanz, V. (Ed), The Shrimp Book 1, 73-88, Nottingham University Press, Nottingham, UK. Hegedus, D., Erlandson, M., Gillott, C., Toprak, U. (2009). New insights into peritrophic matrix synthesis, architecture, and function. Annual Review of Entomology 54, 285-302. Laramore, S.E. (2007). Susceptibility of the peppermint shrimp Lysmata wurdemanni to the white spot syndrome virus. Journal of Shellfish Research 26, 623-627. Lightner, D.V., Redman, R.M., Bell, T.A. (1983). Infectious hypodermal and hematopoietic necrosis, a newly recognized virus disease of penaeid shrimp. Journal of Invertebrate Pathology 42, 62-70. Lin, S.-C., Liou, C.-H., Cheng, J.-H. (2000). The role of the antennal glands in ion and body volume regulation of cannulated Penaeus monodon reared in various salinity conditions. Comparative Biochemistry and Physiology Part A: Molecular & Integrative Physiology 127, 121-129. Mykles, D.L. (2011). Ecdysteroid metabolism in crustaceans. Journal of Steroid Biochemical and Molecular Biology 127, 196-203. Perez, F., Volckaert, F.A.M., Calderon, J. (2005). Pathogenicity of white spot syndrome virus on postlarvae and juveniles of Penaeus (Litopenaeus) vannamei. Aquaculture 250, 586-591. Thuong, K.V., Tuan, V.V., Li, W.F., Sorgeloos, P., Bossier, P., Nauwynck, H. (2016). Per os infectivity of white spot syndrome virus (WSSV) in white-legged shrimp (Litopenaeus vannamei) and role of peritrophic membrane. Veterinary Research 47, 12. Van Thuong, K., Van Tuan, V., Li, W., Sorgeloos, P., Bossier, P., Nauwynck, H. (2016). Effects of acute change in salinity and moulting on the infection of white leg shrimp (Penaeus vannamei)

Chapter 4 µMRI

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with white spot syndrome virus upon immersion challenge. Journal of Fish Diseases 12, 1403-1412. Wheatly, M.G., Gao, Y.P., Nade, M. (2004). Integrative aspects of renal epithelial calcium transport in crayfish: temporal and spatial regulation of PMCA. International Congress Series 1275, 96-103. Young, J. (1959). Morphology of The White Shrimp Penaeus setiferus (Linnaeus 175), Fishery Bulletin 145, 173, The Fish and Wildlife Service, United States of America.

Chapter 5

Nephropore structure and dynamics

during mictio

G.M.A. De Gryse





Chapter 5 Nephropore dynamics and structure

119

5.1. ABSTRACT

Since 1992, when the white spot syndrome virus (WSSV) first emerged in Taiwan, the shrimp

industry has been scrambling to find effective measures to combat this global pandemic.

However, lack of relevant knowledge concerning pathogen entry, makes this task difficult to

accomplish. Fortunately, recent insides put the penaeid shrimp excretory nephrocomplex in

the spotlights as a new potential pathogen entry portal. In Previous chapters, the anatomy

and morphology of this organ appeared to be most favourable for WSSV to use as entry point

and primary replication site. Moreover, considerable involvement in both osmoregulation

and in the moulting process is attributed to the shrimp nephrocomplex. Both are considered

substantial risk factors related to WSSV-infection. However, the underlying mechanisms of

WSSV-entry during the presence of these conditions are still unapprehend. Therefore, in the

current chapter, the defensive structure of the nephropore was investigated using scanning

electron microscopy (SEM) and attempts were made to breach this first line of defence. Using

the SEM information, biodynamic ex vivo models were set up to test the capabilities of theses

defensive mechanisms. Passive pressure from inside the animal directed on the inside of the

nephropore valves could easily open the nephropore to produce an efflux. In contrast, when

pressure was exerted on the outside of the nephropores, no fluid influx or particle entry could

be observed. A final experiment simulated the mictio process, which is a combination of inside

pressure followed by outside pressure. During this simulation, particle entry was observed

using fluorescent beads with sizes corresponding to viruses and bacteria. These experiments

confirmed the defensive competence of the nephropore, but also demonstrated its weakness

during the urination process. This means that conditions during which shrimp urinate more

frequently are to be considered important risk factors for WSSV-outbreaks. These findings fill

an important part of the pathogenesis puzzle and could immediately provide the industry

with useful measures in the battle against WSSV and potentially other important pathogens.

Key words: Penaeus vannamei, nephropore, nephrocomplex, antennal gland, urination,

WSSV, particle entry


120

5.2. INTRODUCTION

The cuticula is a sound barrier protecting shrimp from treats such as dangerous chemicals,

parasites, bacteria, and viruses (Corteel et al., 2009; Corteel and Nauwynck, 2010; Xiong et

al., 2017). The complete external surface of these animals is covered by such a protective

shield. Even the insides of the alimentary tract are covered by either a cuticula or a peritrophic

membrane (Young, 1959; Thuong et al., 2016). When no predispositions are present where

the cuticula is damaged by deficiencies, aggression or chitinases (Keating and Dagbusan,

1984; Sindermann and Center, 1989; Corteel et al., 2009), pathogens must find a different

way to enter shrimp. In Chapter 3, it was established that the lumen of the nephrocomplex is

completely void of cuticular or other chitinous lining. However, the entry of the

nephrocomplex is guarded by the cuticular nephropore (Young, 1959). Chapter 3 and 4 raised

the suspicion of the nephrocomplex to be a major portal of pathogen entry. For pathogens to

enter the nephrocomplex, the protective integrity of the nephropore must therefore first be

compromised. Since the nephrocomplex is an excretory organ in Penaeids and other

Crustaceans (Buranajitpirom et al., 2010), urination must take place at some point. It is the

only moment during which the authors are aware that the nephropore opens.

This chapter aims to investigate the mechanical defence mounted by the nephropore and the

potential dangers of the mictio process for Penaeus vannamei shrimp regarding particle entry

in the nephrocomplex.



Animals

Six SPF P. vannamei shrimp reared in the Artemia Reference Center (Faculty of Bioscience

Engineering, Ghent University) were transferred to the laboratory of Virology (Faculty

Veterinary Medicine, Ghent University). Upon arrival, they were individually housed in

transparent PVC-aquaria of 10L at a constant temperature of 28°C at 35 ppt artificial seawater

with constant aeration. The animals were left to acclimatise for 24h.


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Sample preparation and Image acquisition

Animals were euthanised by cutting of oxygen supply to preserve the external morphology.

Once dead, the shrimp were stripped of the abdomen, mandibulae, maxillae, pereiopodae,

and mandibular palpae, to expose the nephropore. The leftovers where fixed in formol at 4°C

for 4 days.

Based on preservation of the structures of interest, three shrimp where selected. The post

fixation step with osmium tetroxide was skipped due to the dense nature of the exoskeleton.

The sample was dehydrated by submitting it to subsequent washing steps of ultrapure water,

ethanol, and acetone. Next, the sample was washed 3 times 10 minutes in ultrapure water.

Then, it was washed again with subsequently 50% ethanol, 70% ethanol, 94% ethanol, 50%

acetone/50% ethanol (94%), 70% acetone/30% ethanol (94%), and 50% acetone/10% ethanol

(94%). Finally, the sample was left overnight in 100% acetone.

Critical point drying was applied to remove all remaining fluids from the sample by gradually

replacing the acetone by liquid CO2 inside a hyperbaric chamber. The liquid CO2 was finally

turned to gas by exceeding the critical point of CO2.

Next, the sample was mounted on a specimen holder and the surface of the sample was

coated with platina particles through a process called sputtering inside the JFC 1300 Auto Fine

Coater (Jeol Ltd, Zaventem, Belgium) with a pressure of 0.5 mb.

The specimen were placed inside the Jeol JSM 5600 LV Scanning electron microscope (Jeol

Ltd., Zaventem, Belgium) in the Laboratory of Morphology (Faculty of Veterinary Medicine,

Ghent University).

5.3.2. Histology

Animals

Early pre-moult SPF P. vannamei shrimp (MBW 9.13 ± 0.92 g) imported from the Netherlands

and reared in the Artemia Reference Center (ARC, Faculty of Bioscience Engineering, Ghent

University, Belgium) were screened for moulting stage C according to Corteel et al. (2012) by


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inverted light microscopical inspection of the uropods (tail fan) and brought to the Laboratory

of Virology (Faculty of Veterinary Medicine, Ghent University, Belgium). At the Laboratory of

Virology, the shrimp were able to acclimatise for 2 days at 28°C in separate aerated 10 L PVC

tanks at a saltwater concentration of 35 g/L. After this acclimatisation period and recovery

from possible transport related stress, they were euthanised with ice and subsequently

transported to the Laboratory for Light and Electron Microscopy, Department of Morphology

(Faculty of Veterinary Science, Ghent University, Belgium).

Sample preparation and image acquisition

The cephalothoraxes were cut from the abdomen without compromising the hemocoel space

to keep internal configuration as intact as possible. Next, the cephalothoraxes were fixed by

injection of Bouin fixative, a solution of paraformaldehyde and picric acid (85 mL of 2%

paraformaldehyde and 15 mL of saturated picric acid, pH 7.4) and mobile immersion in the

same solution at 37°C for 12 h. Following an overnight washing step in 70 ethanol, the

cephalothoraxes were dewatered with ethanol and xylene in a STP 420D Tissue Processor

(Microm, Prosan, Merelbeke, Belgium) and embedded in paraffin with an EC 350-2 Modular

Tissue Embedding Centre (Microm, Prosan, Merelbeke, Belgium). Ten µm thin slices were

made using a HM360 Rotor Microtome with water bath (Microm, Prosan, Merelbeke,

Belgium) and caught with gelatine coated microscope slide. After drying, the coupes were

stained with a classic hematoxylin-eosine (HE) staining. Finally, cover slips were mounted with

a drop of mounting fluid (DPX, Sigma-Aldrich, St. Louis, USA). The slides were visually

evaluated using an Olympus BX50 light microscope (Aartselaar, Belgium).

5.3.3. Determination of nephropore opening pressure in an ex vivo model

For the analysis of the urinal flow through the nephropore, an ex vivo-model was set up to

determine the pressure needed to open the nephropore valve and let urine flow from inside

the shrimp to the outer world. Shrimp (MBW 15 g ± 1.02 g) of all moulting stages, originating

from the ARC, were euthanized. The antennal appendix of each shrimp was carefully

segregated from the rest of the shrimp body (Fig. 6). All inside tissues of the antennal

appendix were removed with great care under a stereomicroscope (Bausch and Lomb),


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leaving only the exterior shell i.e., the cuticula with the nephropore. Next, the sample was

mounted on a 50 mL plungerless syringe (Terumo Europe N.V., Belgium) such that the

nephropore was situated in the middle of the luer tip opening of the syringe with the inside

of the cuticula facing the lumen of the syringe. A first fixation was achieved with superglue

(Pattex, Henkel, France) and after five minutes, the appendix was embedded into TEC7

(Novatech International, Belgium) without obstructing the nephropore. The construct was left

to dry for three hours (Fig. 1). To measure the pressure needed to evacuate fluid from inside

the shrimp to the outside, the syringe was filled with water until water leaked out of the

nephropore. The height of the water was measured, and the pressure was calculated using

the formula: P = ρ x g x h where P equals pressure (in Pascal), ρ equals the density of the liquid

(1000 kg/m³), g equals the acceleration of gravity (9.81 m/s²), and h equals the leaking height

of the fluid in the syringe. To determine the flow from outside to inside the shrimp, the whole

procedure was repeated with one difference: the nephropore was now not placed on the luer

tip facing up, but facing down, thus simulating possible influx of water in the shrimp.

Fig. 1. Schematic representation of two ex vivo models to test the functionality of the nephropore. Left: ex

vivo model to test the pressure needed to open the nephropore. Right: Ex vivo model to test particle entry during

the mictio process. 1: plastic syringe, 2: luer tip, 3: cuticula with nephropore, 4: TEC7, 5: TEC7 + Bison superglue,

6: Cut base of a cryotube.


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5.3.4. Determination of fluid and particle influx in an ex vivo model after simulated

urination

The base of a plastic cryotube was cut off at 1 cm height and a hole was made in the bottom.

Through this hole, a 1 mL syringe without plunger was introduced and fixed using superglue

(Pattex, Henkel, France). Next, the base of the cryotube was filled with TEC7 (Novatech

International, Belgium) leaving the aperture of the syringe uncovered by the substance. The

glue was left to dry overnight. The antennal appendix, on which the nephropore is situated,

was removed from the shrimp as described above (3.1.1.) and placed on top of the syringe

opening and fixed with superglue, the inside of the cuticula shell facing the luer tip. When

dry, a mixture of TEC7 and a neoprene-based Bison glue (Bison International BV, The

Netherlands) was used to engulf the base of the antennal appendix (coxipodite) ensuring a

watertight seam between the dried TEC7 and the cuticula. The glue was left to dry in 35‰

saline water overnight so the cuticula did not dry out. Next, only the antennal protopodite

was kept while the distal parts (the second antenna (ischiocerite, merocerite, carpocerite, and

flagellum) and the scaphocerite) were removed. The resulting holes were filled with a mixture

of TEC7 and neoprene-based Bison glue to ensure water tightness (50/50). Again, the glue

was left to dry in 35‰ saline water overnight. The entirety was subsequently enrobed with a

layer of superglue, leaving only the nephropore exposed (Fig. 1). Functionality tests using

coloured water and a plunger, proved that the valve-like mechanic of the nephropore was not

altered after this procedure nor was the construct leaking water through the seams.

Afterwards, the syringe was filled with 0.5 mL DNA/RNAse free water and the plunger was

reintroduced. The nephropore was then, submerged in 100mL DNA/RNase free water with

35ppt salt containing 1 µm and 0.2 µm fluorescent beads (Distrilab, The Netherlands) at a

concentration of 105 mL-1 each. The contents of the syringe were quickly pushed into the

solution, simulating urination, followed by a short and quick pull of the plunger, mimicking

the vacuum effect inside the cephalothorax following urine evacuation. The device was

removed from the mixture and rinsed with running DNA/RNAse free water and dried. The

plunger was then pushed again with the nephropore above a microscope slide. A cover slip

was mounted with nail polish and the presence of particles was examined under a confocal

fluorescent microscope (Leica, Germany).


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5.4. RESULTS


The SEM images gave a detailed view on the shrimps nephropore. The nephropore is situated

on the medial side of the antennal coxipodite. The SEM (Fig. 1) revealed the opening of the

nephropore to be a concave, vertical slit. The slit itself is situated on a crater on the medial

side of a spherical protuberantia on the coxipodite. The crater faces caudally. The slit of the

nephropore is formed by two sides of the bottom of the crater, where the caudal side is

superimposed on the rostral side, resembling a check valve-like configuration.

Fig 2. Scanning electron microscopy of the antennal coxipodite with the nephropore. 1: rostral valve and 2:

caudal valve. The white arrowheads point the nephropore split and the black arrowhead points to the rostrum.

5.4.2. Histological examination of the nephropore

On the HE cross-section, the same configuration is visible as observed by the SEM. The

nephropore slit is situated on the bottom of a crater on the protuberantia on the medial side

of the coxipodite. The slit runs vertically and is made up by two superimposing layers of

cuticula. The nephropore is entirely made of exoskeleton of the same sort as the rest of the

external shrimp surface. Furthermore, no smooth muscle layers were found around these

valves nor were there other muscular structures such as sphincter present in the close

proximity of the nephropore slit (Fig. 3).


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Fig 3 HE-staining of the nephropore. A. Localisation of the nephropore (encircled) on the medial side of the

coxipodite, B. Close up of the nephropore slit (black arrow).

5.4.3. Determination of nephropore opening pressure in an ex vivo model

An ex vivo-model was set up to test whether a build-up in urine pressure on the valve is

sufficient for fluid evacuation (simulating the mictio). By dissecting the shrimp and carefully

removing all tissue inside the antennal appendix, the cuticula was mounted on the tip of a 50

mL syringe. A first fixation was performed with superglue and TEC7. Water was poured into

the syringe until drops started to leak out of the nephropore (Fig. 1). The height of the water

level in the syringe (mmH2O) corresponds with a certain hydrostatic pressure, and this was

found to be equal over different moulting stages (2.8 bar). When the nephropore was

reversely mounted on the syringe tip, no leakage was found even after high manual pressure

was exerted (simulating water pressure from outside to inside the shrimp).

5.4.4. Determination of fluid and particle influx in an ex vivo model after simulated

urination.

A second ex vivo model was designed to determine the fluid and particle influx after simulated

mictio (Fig. 1). The nephropore was mounted on the tip of a syringe filled with water. Mictio


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was simulated with the nephropore bathing in a 35‰ salt solution containing green

fluorescent particles (0.2 µm and 1 µm, representing WSSV and V. campbelli, respectively) by

emptying the syringe (pushing the plunger) and abruptly retracting the plunger (creating a

vacuum inside the syringe). After the simulation, particles were found inside the syringe by

confocal microscopy.

5.5. DISCUSSION

This chapter showed through SEM and HE, that the nephropore slit is made up of a rostral

exoskeleton part, superimposed by a caudal exoskeleton part on the bottom of a crater on a

protuberans of the coxipodite. Such a confirmation highly resembles a check valve-like

configuration, which would allow an elastic closing of the aperture. No sphincter-like

structure or other smooth muscles were found around the nephropore, suggesting a passive

opening and closing of the check valve-like structure. Ex vivo experimentation in this chapter,

calculated that when a certain pressure is exerted from inside the nephrocomplex, the check

valve-like structure is allowed to open.

In chapter 3, the muscles associated with the nephrocomplex were studied. Around the

ventral bladder, three important muscles can be found: the coxipodite adductor (CA) and the

small and large scaphocerite abductors (SSA and LSA, Fig 4). Both the CA and the SSA,

originate from the LSA. The LSA muscle attaches in the scaphocerite, to run through the

coxipodite. Just rostrally from the ventral bladder, the SSA and the CA branches split from the

main muscle. The SSA runs lateral from the ventral bladder and the CA runs medially, while

the LSA lays ventrally from the ventral bladder. All three muscles end on the inside of the

exoskeleton, caudally from the ventral bladder. This conformation effectively encloses the

ventral bladder. A coordinated engagement of these three muscles, potentially aided by the

smooth muscle layer in the ventral bladder wall, would result in a higher pressure inside the

ventral bladder and cause the passive opening of the nephropore valves and evacuation of

urine (mictio).


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Fig. 4. Muscles associated with the ventral bladder. VB: ventral bladder; LSA: large scaphocerite abductor; SSA:

small scaphocerite abductor; CA: coxipodite adductor; CGC: compact glandular compartment; VBE: ventral

bladder extension; VBE*: ramifications of the VB; n: nerve; E: exoskeleton.

Another ex vivo set up allowed the demonstration of the structural integrity of this check

valve-like nephropore as high pressure from outside the shrimp on the nephropore did not

allow fluid to be leaked through the nephropore aperture. This confirms the barrier function

exerted by the check valve-like nephropore configuration at rest.

If the nephrocomplex would be a major pathogen entry portal, pathogens should be able to

break this structural integrity of the nephropore check valve-like system. Physical damage

could in theory result in less efficient closing of the valves, but nephropore damage is not

observed to be a necessary feature in WSSV or other infections. Thus, the opening and closing

of the nephropore must be involved in the pathogen entry process. The most apparent reason

for opening the nephropore valves, is during the evacuation of urine from the ventral bladder

of the nephropore to the ambient water (mictio), in which the shrimp is swimming. A final ex

vivo contraption proved that when mictio is simulated, particles are allowed to pass the

nephropore barrier and enter the nephrocomplex. As theorised above, when shrimp urinate,

muscles surrounding the ventral bladder contract, resulting in increased pressure on the

inside of the nephropore. When the pressure reaches a certain threshold, the valves open

and the urine can flow out of the nephrocomplex. During the mictio process, the pressure

inside the ventral bladder will gradually drop because more and more fluid is leaving the

nephrocomplex. Eventually, the pressure inside the shrimp will be equal to the pressure


129

outside the shrimp added to the pressure required to open the valves. At this point, the urine

flow cedes. When the muscles relax, the ventral bladder will expand again and a negative

pressure inside the ventral bladder is created (vacuum). The nephropore will now close again

due to the elastic properties of the exoskeleton (Nishino et al., 1999). The window between

the point where the pressures are equal and the point where the nephropore closes, a proved

influx is possible. We hypothesize now, that this is how pathogens could be able to enter the

nephrocomplex.

According to Young (1959), the SSA and LSA function is to abduct the scaphocerite. Lateral

movement of the scaphocerite is associated with grooming of the cephalon appendices and

a sudden flush from the gill chamber. Given the orientation of the nephropore in the direction

of the gills, the function of the SSA and LSA and position of these muscles around the ventral

bladder, it is plausible that a forceful scaphocerite abduction results in a urine flow towards

the gills. The gills are covered by a lateral, leaf shaped extension of the carapax, forming the

gill chamber (Young, 1959). It is described that the gill chamber possesses epipodites and

exopodites to generate a gentle water flow through the gill chamber. Such a flow could serve

to make oxygen rich water more readily available to the gills, but also to clean the cavity and

to protect it against pathogen colonisation. However, in Penaeid shrimp, these epipodites and

exopodites are strongly reduced and these structures do not suffice on their own to hinder

the building up of sediments trapped in the gill chamber and prevent bacterial and ciliary

overgrowth (Bauer, 1998, 1999). When the abovementioned muscles coordinate and contract

forcefully, a stream of urine could be ejected towards and through the gill chamber. This could

help the shrimp in the process of mechanically cleaning the gill chamber.

It is of course preferable that these ex vivo models would be performed by in vivo

experimentation. However, technologies such as µCT and µMRI are not yet developed enough

to detect such small volumes of contrast fluid. Other experiments using fluorescent beads,

ink or colloid fluids failed because of several reasons. Labelling of viruses with markers would

also be a preferable option to detect particle entry in shrimp, but this technology cannot be

applied to WSSV as no stable cell cultures exist of aquatic invertebrates. Also, the large

genome of the virus (280 to 312 kbp) along with limited knowledge on the functions of viral

proteins, make engineering labelled virus a near impossible feat at this time.


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5.6. CONCLUSIONS

The nephropore resembles a check valve-like structure and is very efficient in guarding the

opening to the nephrocomplex. However, this structural integrity is compromised during the

mictio process when particles the size of viruses and small bacteria are allowed to pass the

nephropore and potentially enter the nephrocomplex.


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5.7. REFERENCES

Bauer, R.T., 1998. Gill-cleaning mechanisms of the crayfish Procambarus clarkii (Astacidea : Cambaridae): experimental testing of setobranch function. Invertebrate Biology 117, 129-143. Bauer, R.T., 1999. Gill-cleaning mechanisms of a dendrobranchiate shrimp, Rimapenaeus similis (Decapoda, Penaeidae): Description and experimental testing of function. Journal of Morphology 242, 125-139. Buranajitpirom, D., Asuvapongpatana, S., Weerachatyanukul, W., Wongprasert, K., Namwong, W., Poltana, P., Withyachumnarnkul, B., 2010. Adaptation of the black tiger shrimp, Penaeus monodon, to different salinities through an excretory function of the antennal gland. Cell and Tissue Research 340, 481-489. Corteel, M., Dantas-Lima, J.J., Wille, M., Alday-Sanz, V., Pensaert, M.B., Sorgeloos, P., Nauwynck, H., 2009. Molt stage and cuticle damage influence white spot syndrome virus immersion infection in penaeid shrimp. Veterinary Microbiology 137, 209-216. Corteel, M., Nauwynck, H.J., 2010. The integument of shrimp, in: Alday-Sanz, V. (Ed), The Shrimp Book 1, 73-88, Nottingham University Press, Nottingham, UK. Keating, K.I., Dagbusan, B.C., 1984. Effect of selenium deficiency on cuticle integrity in the Cladocera (Crustacea). Proceedings of the National Academy of Sciences 81, 3433-3437. Nishino, T., Matsui, R., Nakamae, K., 1999. Elastic modulus of the crystalline regions of chitin and chitosan. Journal of Polymer Science Part B: Polymer Physics 37, 1191-1196. Sindermann, C.J., Center, N.F., 1989. The Shell Disease Syndrome in Marine Crustaceans - A conceptual approach Journal of Shellfish Research 10, 491-494. Thuong, K.V., Tuan, V.V., Li, W.F., Sorgeloos, P., Bossier, P., Nauwynck, H., 2016. Per os infectivity of white spot syndrome virus (WSSV) in white-legged shrimp (Litopenaeus vannamei) and role of peritrophic membrane. Veterinary Research 47, 12. Xiong, H., Pears, C., Woollard, A., 2017. An enhanced C. elegans based platform for toxicity assessment. Scientific Reports 7, 9839. Young, J., 1959. Morphology of The White Shrimp Penaeus setiferus (Linnaeus 175), Fishery Bulletin 145. The Fish and Wildlife Service, United States of America, p. 173.

Chapter 6

Artificial and natural infection of the

nephrocomplex

G.M.A. De Gryse





Chapter 6 Infection of the nephrocomplex

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6.1. ABSTRACT

Aquaculture is often seen as a future sustainable food production source to provide the world

with enough quality protein to sustain the global population. The shrimp industry is

considered to play a major part in this narrative. However, to fulfil this prophecy, losses due

to pathogens must be substantially reduced. The white spot syndrome virus (WSSV) is

arguably the most notorious of shrimp diseases, causing socioeconomic problems worldwide.

Also, bacteria like Vibrio cause a lot of economical loss. Currently, efficient prophylactic and

preventive measures are lacking. Confoundment regarding the entry portal of the pathogens

is to be considered the main culprit. Findings in previous chapter made it clear that the

nephrocomplex is most likely the entry portal of WSSV, researchers have long been searching

for. In this chapter, the main objective is to demonstrate the sensitivity of the nephrocomplex

for WSSV and Vibrio infection with intrabladder inoculation, bypassing the nephropore

defences. Next, Intrabladder inoculation is compared to peroral inoculation and

intramuscular injection. Finally, salinity shock, a known risk factor for WSSV, was applied to

establish an infection via more natural circumstances without artificial inoculation. Samples

were taken from the shrimp’s urine and hemolymph every 12 hours and checked with qPCR

for viral copies. Results showed that WSSV indeed infects the shrimp via the nephropore and

nephrocomplex, where primary replication takes place before a general infection is

established. These findings are a major scientific breakthrough in the field and pave the way

for oriented disease control in shrimp aquaculture industry.

Key words: Penaeus vannamei, Nephrocomplex, antennal gland, WSSV, infection


136

6.2. INTRODUCTION

White spot syndrome virus (WSSV) is regarded as a major cause of production losses within

crustacean aquaculture, which is estimated at 10% or several hundred million US$ annually

(Escobedo-Bonilla et al., 2008; Walker and Mohan, 2009; Pradeep et al., 2012). This virus is of

particular concern to leading scientists because it hinders the future global food supply as

aquaculture is one of the most important food sources to meet the increasing demand of a

growing global population. Shrimp cultivists face complete wipe-out when WSSV infects naïve

populations causing serious sociological drama. Production losses due to chronically infected

shrimp ponds can also not be disregarded (Stentiford et al., 2012). Viruses are not the only

pathogens known to cause serious damage in shrimp industry; bacterial infections such as

vibriosis cause an estimated 20% production loss annually (Hong et al., 2016).

Three routes have been proposed in the past for the transmission of WSSV: (i) through feeding

of infected shrimp (Chang et al., 1996; Zhang et al., 2006), (ii) waterborne, by exposure to

water contaminated with WSSV (Flegel et al., 1997; Tuyen et al., 2014; Domínguez-Borbor et

al., 2019) and (iii) transovarially (Lo et al., 1997). The gut (per os) has been speculated to be

the primary target organ that allows WSSV to enter shrimp. However, the infection efficiency

which can be reproduced via this route is extremely low and remains controversial (Chang et

al., 1996; Wang et al., 1998; Thuong et al., 2016a). Consequently, there is still a lot of debate

on the exact portal of entry (Corteel et al., 2009a; Sánchez-Paz et al., 2012; Dantas-Lima,

2013; Domínguez-Borbor et al., 2019). The gut, along with almost every other structure, is

completely covered by a non-penetrable layer of cuticula or a peritrophic membrane (Bell and

Lightner, 1988; Thuong et al., 2016a). Recent research showed that the cuticula and

peritrophic membrane indeed serve as firm barriers against WSSV (Corteel et al., 2009a;

Thuong et al., 2016a). Therefore, the question was raised on how WSSV (and other pathogens

such as Vibrio) can overcome this barrier. In our search for structures that are connected to

the outside world but have no cuticle-lined lumen, we came across the antennal gland. In the

previous chapters, the potential of the antennal gland (now called nephrocomplex) was

positively assessed as a portal of entry for pathogens. The nephrocomplex has been reported

by other researchers to be amongst the very first organs to become infected using

immunohistochemistry (Rodríguez et al., 2009). Furthermore, Thuong et al. (2016b) showed

increased susceptibility to WSSV infection upon sudden change of salinity. Hemolymph


137

filtration and osmoregulation is the primary function of the nephrocomplex (Freire et al.,

2008; Buranajitpirom et al., 2010). Therefore, these findings further strengthened the

suspicion in the direction of the nephrocomplex as a major portal of pathogen entry and

primary replication site.

Antecedently, some pathogens have been retrieved from the nephrocomplex and analogue

organs of Crustacea. Thrupp et al. (2013) for example, found a parasitic infection in the

antennal gland of juvenile crabs. In the same manner, Pina et al. (2007) isolated Cercaria

sevillana cysts from the green crab’s antennal gland whereas other researchers found a

herpesvirus-like particle through electron microscopical examination of the antennal gland

(Ryazanova et al., 2015). Thus, the presence of pathogens inside the antennal gland or

nephrocomplex is not a new given. However, to this day, no evidence has been provided that

the nephrocomplex serves as an entry portal for pathogens and can act as a primary

replication site. The current study aims to investigate the role of the nephrocomplex as a

possible entry route and primary replication site of WSSV and Vibrio species.


6.3.1. artificial infection

Experimental animals

Specific pathogen-free (SPF) penaeid shrimp, Penaeus vannamei, with a mean body weight of

2.8 g were reared in the Aquaculture & Artemia Reference Center (ARC), Ghent University,

Belgium. Shrimp were cultured in a bio-filter circulation system, fed with pelleted feed at a

rate of 5% of their mean body weight per day. Temperature and salinity of the culture system

were maintained at 27 ± 1oC and 35 ± 1 g L-1. Total ammonia and nitrite were controlled to be

lower than 0.5 and 0.15 mg L-1, respectively. For the experiments, shrimp in early pre-moult

were screened (Corteel et al., 2009b) and transported to the Laboratory of Virology, Faculty

of Veterinary Medicine, Ghent University.


138

WSSV production

The WSSV Thai-1 used in the present study was collected from naturally infected Penaeus

monodon in Thailand in 1996 and passaged in crayfish Pacifastacus leniusculus

(Jiravanichpaisal et al., 2001). Crayfish gill suspension containing WSSV Thai-1 was kindly

donated by K. Söderhäll (Uppsala University, Sweden). The virus was amplified in SPF P.

vannamei juveniles to produce a virus stock. The median infectious titer of the stock was 106.6

Shrimp Infectious Dose50 (SID50) mL-1 i.e., 106.6 times the experimentally determined dose of

virus needed to infect 50% of shrimp, as determined by in vivo intramuscular titration in SPF

P. vannamei (17). A 10-2 dilution of this stock was made in phosphate-buffered saline (PBS),

pH 7.4, and injected intramuscularly into SPF P. vannamei juveniles to amplify the virus. Then,

moribund shrimp were collected and confirmed to be WSSV positive by indirect

immunofluorescence (IIF). Thawed shrimp without shell, hepatopancreas and gut were

chopped, suspended in PBS at a ratio of 1:3, homogenized at 5000 rpm for one minute using

an IKA T 25 digital Ultra-turrax and centrifuged at 5000 g for 20 minutes (4oC). Supernatant

was collected, filtered (0.45 µm) and aliquoted for storage at -70oC. All manipulations were

done inside a laminar flow cabinet under sterile and precooled conditions.

Vibrio campbellii production

Rifampicin-resistant bacteria (Vibrio campbellii: LMG21363) that are pathogenic for penaeid

shrimp were obtained from the ARC, Ghent University, Belgium. From the original stock, 20

µL of bacterial suspension was inoculated in 20 mL Marine Broth (MB) 2216 (Difco

Laboratories, USA) containing 100 mg L-1 of rifampicin for 12 h at 27°C in a shaker at 90

rotations per minute (rpm). Then, the bacteria were subcultured under the same conditions

for 14 h. Afterwards, the suspension was washed and centrifuged three times at 2000 g for

10 minutes. Finally, an estimated concentration of 1010 cfu mL-1 of bacterial suspension (stock)

was made by determining the optical density using spectrophotometry at an absorbance of

600 nm (OD600). An optical density value (OD600) of 1.0 corresponds to 1.2 x 109 cells mL-1

(McFarland standard).


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Infectivity of WSSV stock in P. vannamei by different routes of inoculation

The aim of this study was to compare the infectivity of WSSV stock in shrimp using different

inoculation routes: intramuscular injection, peroral, and intrabladder inoculation. In the

experiment, early pre-moult P. vannamei (25.4 ± 3.3 g) were collected and acclimated

individually for 24 h in 10-liter tanks. Then, 15 shrimp were injected intramuscularly with 5 µL

of a 10-fold serial dilution (10-5, 10-6 and 10-7) of the WSSV stock, as prepared in section 5.1.2,

per animal (5 animals per dilution). Twenty shrimp were inoculated perorally with 50 µL of a

tenfold serial dilution (100 to 10-3) of the same WSSV stock per animal (5 animals per dilution).

A further 20 shrimp in the third group were inoculated into the bladder of the nephrocomplex

with 5 µL of a tenfold serial dilution (10-3 to 10-6) of the same WSSV stock per animal (5 animals

per dilution). The inoculation procedures were fully standardized. Briefly, intramuscular

injection was performed with a 25-gauge needle (Terumo) mounted on an accurate syringe

(Model 1710 LT SYR, 100 µL, Hamilton Bonaduz) filled with 5 µL of WSSV suspension. For oral

inoculation, shrimp wrapped in tissue paper were placed ventral side up under a

stereomicroscope. The tip (2mm) of a 0.64 x 19 mm-26G Surflo-W catheter (Terumo)

mounted on a 100 µL Hamilton Bonaduz, filled with 50 µL of a WSSV suspension, was

introduced into the oral cavity and the inoculum was delivered into the lumen of the foregut.

Based on previously gathered morphological data, an intrabladder inoculation was

developed. First, urine was removed from the bladder after gently introducing the tip (0.5

mm) of a 0.64 x 19 mm-26G Surflo-W catheter (Terumo) in the nephropores. The catheter

was kept stable for a few seconds until the urine fully filled the catheter. Afterwards, the

catheter was removed and replaced with a new 0.64 x 19 mm-26G Surflo-W catheter

(Terumo) connected with a 100 µL Hamilton Bonaduz syringe, filled with 5 µL of a WSSV

suspension. The WSSV suspension was inoculated by exerting a gentle pressure on the

plunger of the syringe. After inoculation, shrimp were housed individually and kept for 5 days.

The intrabladder inoculation was priorly validated on deceased shrimp using 30% iodixanol

solution and control with x-ray microcomputed tomography (µCT) at the Infinity lab (Ghent

University hospital, Belgium) at several volumes. Cephalothoraxes of dead and moribund

shrimp were collected every 12 h. The experiment was terminated at 120 hpi. At the latter

time point all surviving shrimp were euthanized. Samples of dead, moribund, and euthanized


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shrimp were processed for detection of WSSV-infected cells by IIF. The experiment was

performed in triplicate.

Lethality of Vibrio campbellii in P. vannamei by different inoculation routes

From a bacterial stock (1010 cfu mL-1), serial dilutions (10-1, 10-2, 10-3 and 10-4) were made in

filtered, autoclaved seawater (FASW). Then, these dilutions were used to determine the LD50

mL-1 in P. vannamei by intramuscular injection, peroral, and intrabladder inoculations. Fifteen

shrimp were injected with 100 µL of a 10-fold serial dilution (10-2, 10-3, and 10-4) of the Vibrio

stock, as produced in section 5.1.3, per animal (5 animals per dilution). Fifteen shrimp were

inoculated perorally with 100 µL of a tenfold serial dilution (100, 10-1 and 10-2) of the same

Vibrio stock per animal (5 animals per dilution). Fifteen shrimp in the third group were

inoculated intrabladder with 10 µL of a tenfold serial dilution (100, 10-1, and 10-2) of the same

Vibrio stock per animal (5 animals per dilution). After inoculation, shrimp were housed

individually and kept for 5 days. Dead and moribund shrimp were collected every 6 h and the

experiment was terminated at 120 hpi. At the latter time point, all surviving shrimp were

euthanized. For counting the bacterial density in dead, moribund, and euthanized shrimp,

shrimp were washed once with 70% alcohol and twice with FASW. Then, the whole shrimp

were homogenized in FASW at a ratio of 1:5 (100) and diluted (10-1, 10-2 and 10-3) in FASW.

One hundred microliters of each dilution were plated on marine agar containing 100 mg L-1

rifampicin (MAR). The plates were incubated at 28oC for 24 h, then counting was conducted

with each plate containing between 30 and 300 colonies.

Replication of WSSV in P. vannamei after intramuscular and intrabladder inoculation

Early pre-moult SPF-shrimp (25.2 ± 3.8 g), acclimated for 1 day at 27oC, were divided into

three groups. Animals were injected intramuscularly with 10 µL containing 105.5 SID50 of WSSV

at the junction between the fourth and fifth abdominal segment. Animals were inoculated

intrabladder with 10 µL containing 105.5 SID50 of WSSV. Animals were mock inoculated by

intramuscular injection with 10 µL of PBS and used as controls. After inoculation, blood, and

cephalothoraxes of five shrimp were sampled at 0, 6, 12, 18, 24, 36 and 48 hpi. For the

determination of WSSV-infected cells in hemolymph, 500 µL of hemolymph were collected


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with anticoagulant buffer at a ratio of 1:1. Hemolymph was further diluted 1:1 in PBS. Then,

100 µL of the diluted hemolymph was cytospinned. The cytospins were centrifuged at 300 g

for four minutes, and then fixed for ten minutes in methanol. WSSV viral antigens were then

detected by IIF. For the determination of WSSV-infected cells in tissues, the cephalothoraxes

were cross-sectioned (20 μm) at the site of the nephrocomplex, hepatopancreas, gills,

lymphoid organs, heart, and hematopoietic tissues. Then, the sections were fixed in methanol

for ten minutes and WSSV viral antigens were detected by IIF. WSSV-infected cells in the gills,

hepatopancreas, lymphoid organ, coelomosac, heart and hematopoietic tissues were

counted in six randomly selected fields and expressed as the number of WSSV-infected cells

per square millimetre (cells mm-2). WSSV-infected and uninfected cells were counted

randomly in six fields of hemocytes, bladder epithelial cells and cuticular epithelial cells of the

body, head, and hindgut and expressed as a percentage of infected cells (%).

Detection of WSSV viral antigens by indirect immunofluorescence

Viral antigen positive cells in WSSV-infected shrimp were detected by IIF as described by

Escobedo-Bonilla et al. (2006). Briefly, cephalothoraxes of dead, moribund, and euthanized

shrimp were collected, embedded in 2% methylcellulose and frozen at -20oC. Cryosections (6

µm) were made and fixed for ten minutes in 100% methanol at -20oC. The sections were

washed three times in PBS (5 min. each) and incubated in 200 µL of monoclonal antibody w29

(directed against WSSV viral protein VP28 1:100 in PBS) at 37°C for 1 h. Then, samples were

washed three times in PBS (5 min. each), incubated with fluorescein isothiocyanate (FITC)-

labelled goat anti-mouse IgG (F-2761, Molecular Probes, The Netherlands) for 1 h at 37oC.

Finally, the samples were washed twice with PBS, rinsed once in deionized water, and

mounted with a solution of glycerine and 1,4-diaza-bicyclo-octane (DABCO) (ACROS Organics,

USA). Sections were analysed with a confocal fluorescence microscope.

Statistical analysis

Determination of viral infectivity (SID50) and bacterial lethal titers (LD50) was done based on

the method of Reed and Muench (1938).


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6.3.2. Natural infection using salinity shock immersion

Twenty-five shrimp of ± 35 g from the ARC were screened for moulting stage C and brought

to the Laboratory of Virology, Ghent University. After one week of acclimatization, 5 animals

were transferred to a single 10 L PVC tank with a partition in the middle (Van Thuong et al.,

2016). In the first half of the tank, 1 L of artificial seawater (35‰ salinity) was added and in

the other 1 L of 5‰ artificial seawater. The shrimp started the experiment in the 35‰ half

and after one day of acclimatization, the animals were gently flipped to the 5‰ part of the

tank. At the time of the flipping, 105.5 SID50 mL-1 (1 L) of WSSV Thai-1 was present in this 5‰

half of the tank. After five hours, the shrimp were gently flipped back to the 35‰ half of the

tank. Every 12 hours, urine and hemolymph were collected. Urine sampling was performed

with a similar protocol of intrabladder inoculation; the tip (0.5 mm) of a 0.64 x 19 mm-26G

Surflo-W catheter (Terumo) connected to a 1 mL syringe was introduced between the two

valves of the nephropore. As soon as urine started to appear in the catheter due to capillary

flow, a gentle vacuum was created using the syringe, extracting as much urine as possible

without exposing the shrimp with unnecessary stress. Hemolymph was extracted using a

novel method involving the introduction of a needle in the hemolymph sinus next to the

heart. Next, DNA extraction (cador pathogen kit, Quiagen) and subsequently qPCR was

performed on both urine and hemolymph samples (Li et al., 2015). The experiment was

performed in triplicate. Two controls of each five shrimp were simultaneously performed: a

control with a drop in salinity but without WSSV and a control without a drop in salinity but

with WSSV. Both control groups were sampled (urine and hemolymph) and tested with qPCR.

6.4. RESULTS

6.4.1. Validation of intrabladder inoculation

For the validation of the intrabladder inoculation method, the shrimp where first inoculated

with a 30% iodixanol solution and put under µCT. First, urine was removed from the bladder

after gently introducing the tip (0.5 mm) of a 0.64 x 19 mm-26G Surflo-W catheter (Terumo)

in the nephropores. The catheter was kept stable for a few seconds until the urine fully filled

the catheter. Afterwards, the catheter was removed and replaced with a new 0.64 x 19 mm-


143

26G Surflo-W catheter (Terumo) connected with a 100 µL Hamilton Bonaduz syringe, filled

with variable volume of iodixanol 30%. The solution was inoculated by exerting a gentle

pressure on the plunger of the syringe. After inoculation, shrimp were examined under µCT.

For volumes up to 100µl, the iodixanol generated a contrast-rich shape corresponding to the

outlines of the ventral bladder as determined by HE and µMRI in previous chapters. When

higher volumes were introduced, the iodixanol signal was retrieved all over the hemocoel

(Fig. 1).

a. b.

Fig. 1. Micro computational tomography (µCT) scan of contrast fluid upon intra-bladder inoculation. a. Urinary

bladder intact upon inoculation of 100µl of 30% Iodixanol (arrow), b. Urinary bladder disrupted upon inoculation

of 120µl of 30% iodixanol, the contrast fluid can be seen throughout the hemocoel.

6.4.2. Artificial infection of the nephrocomplex

Different inoculation routes were compared. For a given WSSV Thai-1 stock, an infectious titer

of 108.67, 108.67, and 108.80 shrimp infectious dose 50% end point (SID50) mL-1 (x ̅= 108.71 SID50

mL-1) was achieved after intramuscular inoculation; 101.14, 101.30, and 101.30 SID50 mL-1 (x ̅=

101.25 SID50 mL-1) upon peroral inoculation, and 106.97, 106.60, and 107.30 SID50 mL-1 (x ̅= 106.96

SID50 mL-1) after intrabladder inoculation. All dead and moribund shrimp were WSSV positive,

while surviving shrimp were WSSV negative (Table 1). Taken together, this means that

compared with the intramuscular route, only 56 times more infectious virus is needed to

infect shrimp via intrabladder inoculation, whereas 28.8 x 106 times more virus is necessary

to infect shrimp via oral inoculation (Table 1).


144

Exp. route Dilution

of WSSV stock

N Number of dead animals at …. hpi N

IIF positive Infectivity titer

(SID50 ml-1) 24 36 48 60 72 84 96 120 Total

1 im 10-5 5 3 1 1 5 5 108.67 10-6 5 2 2 4 4 10-7 5 0 0

po 100 5 2 2 2 101.14 10-1 5 0 0 10-2 5 0 0 10-3 5 0 0

ib 10-3 5 2 2 1 5 5 106.97 10-4 5 1 2 1 4 4 10-5 5 1 1 2 2 10-6 5 0 0

2 im 10-5 5 3 2 5 5 108.67 10-6 5 1 2 1 4 4 10-7 5 0 0

po 100 5 1 1 2 2 101.30 10-1 5 1 1 1 10-2 5 0 0 10-3 5 0 0

ib 10-3 5 3 2 5 5 106.60 10-4 5 1 1 1 3 3 10-5 5 1 1 1 10-6 5 0 0

3 im 10-5 5 3 1 1 5 5 108.80 10-6 5 2 1 1 1 5 5 10-7 5 0 0

po 100 5 2 2 2 101.30 10-1 5 1 1 1 10-2 5 0 0 10-3 5 0 0

ib 10-3 5 2 2 1 5 5 107.30 10-4 5 2 1 1 4 4 10-5 5 2 1 3 3 10-6 5 0 0

Table 1. Infectious virus titers of a WSSV stock upon intramuscular (im), peroral (po) and intrabladder (ib) inoculation in P. vannamei


145

For a given V. campbellii stock, a similar experiment was performed (Table 2): 104.16, 104.37,

and 104.16 lethal dose with 50% end point (LD50) mL-1 (x ̅ = 104.23 LD50 mL-1) was found on

intramuscular inoculation and 102.50, 102.50, and 102.32 LD50 mL-1 (x ̅ = 102.44 LD50 mL-1) on

intrabladder inoculation. It was not possible to kill shrimp using the peroral route.

Determination of bacteria by plate counting indicated that all dead and moribund shrimp

were V. campbellii positive and contained high densities of V. campbellii (3.8 ± 1.0 x 105 colony

forming units (cfu) mL-1 of homogenate), whereas surviving shrimp showed a clearing

mechanism to eliminate bacteria from their body (2.4 ± 3.4 x 102 cfu mL-1 of homogenate).

Taking all results together, these findings demonstrate that, compared with the intramuscular

route, only 62 times more V. campbellii is required to kill shrimp via intrabladder inoculation

whereas it was not possible to kill shrimp by the peroral route with up to 109.0 V. campbellii

(Table 2).


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Table 2 Lethality titers of Vibrio campbellii upon intramuscular (im), peroral (po) and intrabladder inoculation in P. vannamei.

Experiment Inoculation route

Dilution of Vibrio N

Number of dead animals at different time points Mortality

(%) Lethal titer (LD50 ml-1) 0 6 12 18 24 30 36 120 Total

1 im 10-2 5 5 5 100 104.16 10-3 5 1 2 3 60

10-4 5 0 0 po 100 5 0 0 - 10-1 5 0 0

10-2 5 0 0 intrabladder 100 5 2 1 3 60 102.49 10-1 5 2 2 40 10-2 5 0 0

2 im 10-2 5 3 2 5 100 104.37 10-3 5 2 2 4 80

10-4 5 0 0 po 100 5 0 0 -

10-1 5 0 0 10-2 5 0 0

intrabladder 100 5 2 1 3 60 102.49 10-1 5 2 2 40 10-2 5 0 0

3 im 10-2 5 5 5 100 104.16

10-3 5 2 1 3 60 10-4 5 0 0 po 100 5 0 0 -

10-1 5 0 0 10-2 5 0 0

intrabladder 100 5 2 1 3 60 102.32 10-1 5 1 1 20 10-2 5 0 0


147

Next, the pathogenesis of a WSSV infection in this organ was examined after intrabladder

inoculation. Intrabladderly and intramuscularly inoculated animals were compared (See Fig.

2). In the intramuscular inoculation group, the first WSSV-infected cells were observed at 18

hours post inoculation (hpi) in all investigated tissues: ventral urinary bladder, compact

glandular compartment (coelomosac and labyrinth), heart, gills, lymphoid organs,

hepatopancreas, hematopoietic tissues, and cuticular epithelium of head, body, and hindgut.

WSSV positive cells in hemolymph were detected at 24 hpi. Upon intrabladder inoculation,

WSSV-infected cells were only detected in the epithelial cells of the bladder at 18 hpi. From

24 hpi onwards, WSSV positive cells were seen in the other investigated tissues and in some

cells in the hemolymph. With both inoculation routes, only a small proportion (<2%) of cells

in the hemolymph were WSSV positive. WSSV-infected cell numbers of all investigated tissues

were low at 24 hpi. The number of WSSV-infected cells then increased rapidly and reached

high levels after 36 hpi. The highest count of WSSV-infected cells was found in the

coelomosac, labyrinth, gills, and the cuticular epithelial cells. The number of WSSV-infected

cells observed in the intramuscular inoculated shrimp was higher than observed in the intra

bladder inoculated animals in all investigated tissues

Fig. 2. (Next page) Photomicrographs (PM) of nephrocomplex and surrounding areas of intramuscularly and

intrabladder WSSV-inoculated shrimp sampled at 18 hpi (a, b, c, d) and 24 hpi (e, f, g, h). Left panel: WSSV-

infected cells were detected by IIF using a WSSV VP28-specific mouse monoclonal antibody and an FITC-

conjugated goat anti-mouse IgG (green, see arrows); nuclei were visualized with Hoechst (blue). Right panel: for

a correct orientation, cryosections were stained using Diff-Quick staining. PM a & b show the first WSSV-infected

epithelial cells in the ventral bladder (VB) at 18 h after intrabladder inoculation; 6 hours later (e & f), WSSV

positive cells were found in the ventral bladder, compact glandular compartment (CP) and muscles surrounding

the nephrocomplex. PM c, d, g, h show WSSV-infected epithelial cells in the ventral bladder, compact glandular

part, and surrounding areas of the nephrocomplex at 18 and 24 h after intramuscular inoculation. AM1: adductor

muscle of first antennal segment, AM4: adductor muscle of fourth antennal segment, SA: scaphocerite adductor

muscle. Bar = 200 µm

.


148

.


149

6.4.3. Natural infection of the nephrocomplex

When shrimp were challenged with WSSV during a salinity shock, the virus could be detected

with a quantitative polymerase chain reaction (qPCR) in the urine 12 h post infection (93% of

all challenged animals), while the hemolymph mostly stayed negative (73%). Only after 24

hours, the virus could be detected in the hemolymph of all shrimp (100%). Increasing amounts

of WSSV copies were found in the urine and hemolymph at later time points. All shrimp were

dead 48 hpi. Shrimp of control group 1 (drop in salinity; no WSSV) and of control group 2 (no

salinity drop; with WSSV) remained completely virus negative and all shrimp survived.

Fig.3. Graph representing the results of the viral salinity shock immersion challenge.

6.5. DISCUSSION

In this chapter, the hypothesis of the nephrocomplex as a major pathogen entry portal

candidate, was confirmed by intrabladder inoculation of shrimp with WSSV and V. campbellii,

both leading to morbidity and mortality. We demonstrated that these pathogens could easily

infect shrimp through inoculation of the nephropore (intrabladder). The data shows that


150

WSSV and Vibrio infect shrimp more efficiently via the nephrocomplex (intrabladder).

Compared to intramuscular inoculation, only 56 times more infectious WSSV is needed to

infect shrimp via intra bladder inoculation, while 28.8 x 106 times more virus is necessary to

infect shrimp via oral inoculation. For Vibrio, similar results were obtained (62 and >109.0 times

more, respectively). These data show that the intrabladder infection is almost as efficient as

the intramuscular inoculation, which bypasses all natural defence barriers such as

nephropore valves, cuticula, and peritrophic membrane. In contrast, the peroral route was

significantly less efficient. This was in accordance with the findings of Thuong et al. (2016a).

In the pathogenesis experiment, we clearly demonstrated that, when infected via the

nephropore, WSSV first replicates in the epithelial cells of the bladder at 18 hpi and

afterwards spreads all over the body. This proves the possibility and efficiency of viral spread

from the tissues of the nephrocomplex into the hemocoel where the virus can infect all other

susceptible organs.

Intrabladder inoculation proved to be successful in infecting shrimp with pathogens.

Nevertheless, introducing the pathogens directly into the target organ, thereby avoiding

natural barriers, is very artificial. Therefore, more realistic proof is required. A drop in salinity

has been proven to facilitate WSSV infection (Thuong et al., 2016b). We found, that during

such conditions, shrimp urine tested WSSV qPCR positive before hemolymph did. This is a

strong indication that WSSV first enters the urine (and thus the nephrocomplex) before the

hemolymph and consequently all other organs are reached. After 24 hpi and further on, the

presence of increasing amounts of WSSV copies in urine further supports the

nephrocomplex’s role as primary replication site.

6.6. CONCLUSIONS

Infection of WSSV and Vibrio via the nephrocomplex is a very efficient way to disease shrimp,

much more efficient than e.g., via the per oral route, which has been considered the major

entry portal of WSSV. However, this study demonstrated with both artificial and natural

infection protocols, that the shrimp nephrocomplex is to be considered the major entry portal

of WSSV and even other pathogens.


151

6.7. REFERENCES

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Chapter 7

General Discussion

G.M.A. De Gryse



Chapter 7 Discussion

157

7.1. THE SHRIMP’S NEPHROCOMPLEX (ANTENNAL GLAND) IS MUCH MORE COMPLEX THAN

PREVIOUSLY ASSUMED.

Prior to this thesis, the nephrocomplex was known to exist of a central coelomosac

responsible for hemolymph filtration, surrounded by the anastomosing tubuli of the labyrinth

and ending via a urinary bladder in a nephropore, which connects to the outer world (Xiaoyun

et al., 2003). Young (1959) described colorations above the supra-esophageal ganglion when

dye was injected through the nephropores of Penaeus setiferus. Several other publications

make notice of tubular structures sporadically spread around the hemocoel and even making

functional connections with the lymphoid organ (e.g. Duangsuwan et al., 2008; Rusaini and

Owens, 2010). The first and only real attempt to map the anatomy and morphology of the

nephrocomplex in Penaeus japonicus was performed by two under-the-radar Japanese

studies, which made a clay model based on histological sections (Nakamura and Nishigaki,

1991; Nakamura and Nakashima, 1992). Although a very meritorious attempt (they more or

less managed to describe some bladders), much of the organ structure was mislabelled as

coelomosac and the black-and-white photos of the clay model where very unintelligible.

The modern 3D-reconstruction software AMIRA allowed the authors in chapter 3 to remap

the anatomy and morphology of the shrimp’s excretory gland. The central coelomosac,

surrounded by the tubuli of the labyrinth (collectively named as the compact glandular

compartment), ending in a ventral urinary bladder was confirmed. Besides, several extensions

to both the rostral direction and caudal directions were observed. When the tubuli of the

labyrinth extend rostrally to form the rostral compartment, the epithelium does not change

much when compared to that of the labyrinth. The rostral compartment tubuli fill up the

entire hemocoel of the cephalon, surrounding the supra-esophageal ganglion and making

some connections to the antennal artery, which earlier penetrated the compact glandular

compartment. Finally, the tubuli will return to the labyrinth before ending in the ventral

urinary bladder. This conformation of the rostral compartment of the excretory organ of

shrimp resembles the loops of Henle as found in the mammalian kidney (Morgan and Berliner,

1968). Given the described functions of the labyrinth from which the tubuli of the rostral

compartment originate, and its similar cellular morphology, it is quite possible that the

function of this rostral compartment is surface enlargement for modulation of the filtrate,

again much like the loop of Henle. Another implication of the surface enlargement could


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signify a role in the osmoregulation of the hemolymph (Lin et al., 2000), ion regulation (Dall

and Smith, 1981; Cheng and Liao, 1986; Vogt and Quinitio, 1994), detoxification of heavy

metals (Doughtie and Rao, 1984; Vogt and Quinitio, 1994; Páez-Osuna and Tron-Mayen,

1996), and perhaps also the (fixed phagocytotic) immune function described to the

nephrocomplex (Doughtie and Rao, 1984; Johnson, 1987; Kondo et al., 2012). More surface

means more chances of interaction between the cells and their targets, and therefore possibly

a higher activity of the nephrocomplex. Further molecular characterisation of these tubuli

should provide further insights into their functions.

Using AMIRA-based 3D-reconstruction, and later confirmed with a µMRI-based 3D-

reconstruction in chapter 4, several bladder-like continuations of the nephrocomplex were

uncovered. They all stem from the most caudal site of the ventral urinary bladder; (i) the

median compartment with a ventral and dorsal lobe covering a major part of the stomach,

and (ii) the lateral compartment with its caudal extensions, reaching as far as the

hepatopancreas. Both bladders, like the rest of the nephrocomplex, lack cuticular lining and

can fill large parts of the hemocoel. Skeletal muscles are associated to the compartments, and

the walls of the bladder compartments sometimes contain a smooth muscular layer. This

hints at a dynamic function, which was confirmed by µMRI in chapter 4 (see further). Several

ramifications of the ventral bladder and other compartments can be found all over the shrimp

cephalothorax. This could also purpose surface enlargement, but its functions are still largely

unknown.

The morphology of the organ is now well described. However, the functions of many

nephrocomplex parts remain unclear, e.g., does the rostral compartment only resemble the

loop of Henle morphologically or also functionally? A lot of the functions of the compact

glandular compartment remain vague and are molecularly poorly described. Most of the

functional knowledge of the antennal gland or nephrocomplex is still extrapolated from other

crustaceans and sometimes even insects. Therefore, functional, and molecular

characterisation studies of the coelomosac, labyrinth and rostral compartment cells is

warranted. For example, it would be interesting to know where Na+/k+-ATPases, besides in

the coelomosac and labyrinth, are present. Also, information on the embryological

development of the nephrocomplex over the different shrimp life stages or ages would be

welcome.


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7.2. THE SHRIMPS NEPHROCOMPLEX IS INVOLVED IN THE MOULTING PROCESS

Moulting is a very impactful event on shrimp. Drastic changes occur in the shrimp’s body

when the metabolism changes from homeostasis to allostasis while the animal prepares to

shed its old cuticle and grows new tissue. With a complexity and an almost unrivalled

extensiveness, it is not unlikely that the nephrocomplex plays a significant role in this process.

In chapter 4 the role of the nephrocomplex in this process was investigated. During the

different moulting stages, the volume of the nephrocomplex bladders evolve. An overview of

the role of the nephrocomplex in moulting is given in the next paragraph.

A first part is played by metabolising the moulting hormone ecdysone from the Y-organ and

excretion of its metabolites. This way, the organ can regulate the balance between the moult

inhibiting hormone and moulting hormone ecdysone, which determines the progression

through the subsequent moulting stages (Chang, 1985; Mykles, 2011). The moulting hormone

ecdysone reaches a peak in stage D1 (early pre-moult), urging the cells to prepare for ecdysis.

Late pre-moult these hormone levels drop significantly (Corteel and Nauwynck, 2010).

Immediately before ecdysis, coordinated actions of muscles associated with the

nephrocomplex, pump extra fluids to the caudal extensions (CE) of the lateral compartments.

The contents of the CE are now at a maximum (Fig 10). This higher volume aids in raising the

inside pressure on the carapax, putting ample pressure on the ligament between the carapax

and the first abdominal segment. When the shrimp contracts its other muscles, the added

pressure by the CE helps in shedding of the exuvium. Immediately after the shedding of the

old cuticula, the shell is still soft. This is the only time during the moulting-cycle, when

absolute volume of the shrimp can be increased. Shortly after the ecdysis, the volume inside

the CE remains high, helping shrimp to increase that volume as much as possible while the

exoskeleton hardens. The hardening of the new, still soft shell happens through the process

of mineralisation. Calcium plays a pivotal role in the mineralisation of the new exoskeleton by

forming calcite (CaCO3) and as an intracellular messenger in the hormonal actions (Ahearn et

al., 2004 and Wheatly et al., 2004). Without doubt, the nephrocomplex fulfils an important

part in regulating the calcium concentrations inside the hemolymph, most probably by the

cells of the labyrinth and by the coelomosac and quite possibly by other parts of the organ as

well (e.g., rostral compartment). However, this remains to be investigated. Next, when the

shrimp progress through the moulting stages (A and B), the volume inside the CE will gradually


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drop, making room for newly grown tissue. During intermoult (C) and early pre-moult (D1),

the volume inside the CE will be at its lowest and the moulting inhibiting hormone will be in

balance with the moulting hormone ecdysone (Corteel and Nauwynck, 2010). Another path

wide open to scientific research progress is to further unravel the complete role of the

nephrocomplex in the moulting process. For instance, the median compartment is most likely

also involved in regulating absolute body volume after ecdysis like the caudal extensions are,

but evidence has not yet been provided. The origin of the water taken up by shrimp is also a

point of discussion. The first option is immediate uptake of ambient water via the nephropore,

exposing the shrimp to microbiological and chemical challenges, after which the water is

transported by coordinated muscle movement to the bladder compartments. The second

option is that the water is taken up via the midgut after drinking as observed in lobsters

(Mykles, 1980). The water could then be transferred to the nephrocomplex, either via the

CGC after which the fluids are pumped by to muscles to the bladder compartments or directly

to the bladder compartments where it is held in place by the continuously contracted

muscles.

µMRI-observation of the volume of the bladders during post-moult as described in chapter 4,

could provide a reliable platform for artificial selection of breeding shrimp based on the size

of the caudal extensions around moulting. Hypothetically, the shrimp with the largest bladder

volume, could gain the most tissue during the growing stages.

7.3. THE SHRIMP’S NEPHROCOMPLEX IS A MAJOR ENTRY PORTAL OF PATHOGEN ENTRY

For shrimp to get diseased, various pathogens must first find a way to enter shrimp and reach

the primary replication site. Prior to this work, no study has ever indubitably demonstrated

an entry portal for WSSV or any other systemic shrimp virus. Up till now, both the entry point

and primary replication site still had to be discovered (Lightner, 2011). Many studies assumed

a peroral entry of WSSV (Chang et al., 1996; Wang et al., 1998; Rajendran et al., 1999),

although with low efficiency (Perez et al., 2005; Laramore, 2007; Corteel et al., 2012). Chapter

3 showed the nephrocomplex to be in pole position as the number one suspect for WSSV

entry. The fact that the nephropore is nothing other than a hole in the exoskeleton, the lack


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of inner cuticular lining and proximity to all other susceptible organs, support this statement.

Also, chapter 4 linked the nephrocomplex to the moulting process, which is in its turn also

linked to WSSV-infection (Corteel et al., 2009; Thuong et al., 2016a).

It is of the utmost evidence that shrimp would be more vulnerable to various pathogens if the

numerous potential entry portals were not so efficiently shielded by first line defences. These

defences are generally mounted by some forms of chitinous structures such as the

exoskeleton, peritrophic membrane or the nephropore valves. These barriers not only make

shrimp hard to infect, but they also impede the search for pathogen entry portals (Corteel et

al., 2009). This cuticular layer is not only protective against physical threats but is also very

effective in fending off other noxes such as pathogens and toxins (Corteel et al. 2009). The

cuticula is made of chitin fibres with protein deposits, reinforced by minerals such as calcium

and magnesium. This combination makes the structure strong and resilient, but also very

dense. Furthermore, significant parts of the exoskeleton (epicuticle, exocuticle) are formed

before the old cuticle is shed, insuring moult-cycle round protection (Corteel and Nauwynck,

2010). This cuticula not only enrobes the entire shrimp’s external body (including eyes and

gills), the foregut and hindgut are also completely covered by a non-permeable cuticle layer

(Young, 1959; Thuong et al., 2016b). In contrast, no cuticle is present to protect the midgut.

Instead, a non-cellular, slimy matrix of chitin and proteins called the peritrophic membrane,

ensures local microbial security (Hegedus et al., 2009). It was experimentally demonstrated

that no particles with a diameter greater than 10 nm are able to pass the peritrophic

membrane (Martin, Rebecca et al. 2006). When first barrier defences were bypassed by

introducing a fine catheter in the nephropore in chapter 6 (intra bladder inoculation), we

demonstrated that this way of infecting is about 28.8 million times more efficient for WSSV

as compared to peroral inoculation and only 56 times less efficient when compared to

intramuscular injection, which bypasses all barriers (not only the cuticula, but also coagulation

and melanisation). These numbers are the result of a combination of a protection by the

presence of a cuticle and peritrophic membrane and the resistance of the endoblastic tubular

epithelial cells lining the inside of the gut against a WSSV infection. In contract the epithelium

of the nephrocomplex is clearly susceptible to this virus and facilitates further systemic viral

invasion. Indeed, IIF showed WSSV replication after intra bladder inoculation, confirming the

potential of the epithelium of the ventral bladder to serve as primary replication site. When


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Vibrio campbellii was inoculated, intrabladder inoculation was one billion times more efficient

than peroral inoculation, demonstrating the broad range of pathogens able to cause disease

when entering the nephrocomplex.

In order for these pathogens to reach the entry sites in a natural way, certain occurrences or

conditions capable of compromising these defences called risk factors, must take place.

Concerning WSSV: shrimp which are in post-moulting stages, endure harsh salinity shocks or

are stocked in high densities are most at risk at contracting the devastating syndrome (Corteel

et al., 2009; Tendencia et al., 2011; Thuong et al., 2016a). In chapter 6, it was demonstrated,

that after exposing shrimp to a WSSV-immersion challenge during a salinity shock (35‰ to

5‰), the virus could be retrieved in the urine of the majority of shrimp before it could be

detected in shrimp hemolymph. This shows that, before the virus reaches the hemolymph,

viral replication happened first in the nephrocomplex, after which it was allowed to spread to

the rest of the shrimp body via the hemolymph. At further time points, it should be noted

that the viral load increased in both urine and hemolymph, indicating viral replication in both

the nephrocomplex (as primary replication site) and the other organs (secondary replication

sites). Replication inside the nephrocomplex also means that immense numbers of WSSV can

be expulsed via the urine and increase the viral load in the surrounding body of water during

already the beginning phases of infection.

In chapter 5, the sturdiness of the nephropore as a check-valve like mechanism under resting

conditions was demonstrated, leaving the question how the nephrocomplex defences can be

compromised. Also in chapter 5, experiments simulating the mictio process, proved particle

entry to be possible during the urination process. When pressure builds up inside the ventral

bladder, possibly aided by muscles surrounding this structure (see chapter 3), the nephropore

will open at a certain pressure threshold (see chapter 5). Next, the urine will be evacuated

from the ventral bladder into the outer water world. The pressure inside the ventral bladder

will gradually drop until the pressure inside the bladder is more or less equal to the pressure

outside, causing a momentary status of equilibrium where no net pressure is exerted on the

valve of the nephropore, causing it to float freely between the closed and open position.

When the muscles surrounding the ventral bladder relax, a vacuum effect will be created

causing the nephropore valve to close. The window of opportunity for pathogens to enter the

nephrocomplex can be found between the equilibrium stage and the reclosing of the


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nephropore valves. The amount of unwanted water influx will be minimal, but as

demonstrated by the intrabladder inoculation in chapter 6, only a small amount of virus is

necessary to cause disease, especially when infection pressures are high. Thus, risk factors

leading to an increase of the mictio frequency should provide WSSV with ample opportunities

to breach the nephropore defences and enter the nephrocomplex to cause disease in shrimp.

Salinity shock as described in chapter 6 is such a condition or risk factor and is a confirmation

of the work of Thuong et al. (2016). When the ambient salinity drops, the shrimp will take up

more water and its cells risk an enduring osmotic shock. As a reaction, the shrimp

nephrocomplex, in charge of the osmotic homeostasis of shrimp, will produce more urine (Lin

et al., 2000). More urine produced will most probably lead to increased urination and thus an

increased number of opportunities for pathogens to enter the shrimp’s body. Increased

stocking density is also linked to WSSV-outbreaks (Tendencia et al. 2011) and can be

interpreted in two ways: more shrimp per m³ water means more infectable and infected

shrimp, which in turn, leads to higher virus titers in the pond water and thus to a higher

infection pressure meaning more virus can enter with every mictio. High stocking density also

leads to increased cannibalism (eating of infected tissue), aggressiveness (cuticle damage,

stress) and increased need to establish social dominance, which comes with a higher urination

frequency in many crustacean species (Breithaupt and Atema, 2000; Katoh et al., 2008).

Moreover, when shrimp endure acute stress situations, shrimp will often urinate (personal

observation). Also, when stocking densities are higher, more aggressiveness is likely to occur.

Potentially, fighting animals could injure each other. It could be hypothesised, that if the

nephropore is damaged during skirmishes, the sealing function diminishes, because the

valves will not fit properly anymore, allowing particle influx. Finally, the ecdysis and post-

moulting stages are also linked to WSSV-infection as well (Corteel et al., 2009; Thuong et al.,

2016a). In chapter 5, it was demonstrated, that prior to ecdysis, the contents of the

nephrocomplex increase drastically. After ecdysis, the volume will gradually decrease inside

the nephrocomplex to make room for newly grown tissue. A decrease in volume, means an

increase in urination post-moulting and thus, again, an increased number of opportunities for

pathogens to enter shrimp. Recent independent research came to the same conclusion as this

thesis, namely that salinity stress facilitates WSSV-entry in the nephrocomplex (Liu et al.,

2021).


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Previously, most researchers believed that shrimp were primarily or only infected with WSSV

perorally. This was based on early reports of feeding WSSV-contaminated feed to shrimp,

leading to WSSV-infected animals. However, to the best of the authors knowledge, no reports

exist demonstrating WSSV entry along the digestive tract. Furthermore, removal of the

peritrophic membrane did not augment shrimp WSSV-susceptibility and high viral titers are

required to attain WSSV infection through feeding (Perez et al., 2005; Laramore, 2007;

Thuong et al., 2016b). The contrast to WSSV entry after immersion via the nephropore could

not be higher: chapter 6 showed intrabladder inoculation to be highly efficient, qPCR- and IIF-

proof of primary WSSV-replication in the bladder epithelium, and particle entry during the

urination process has been demonstrated both artificially and naturally. A peroral inoculation

or a feeding challenge of shrimp could quite effectively lead to a de facto immersion

challenge. Shrimp eat very violently, shredding big chunks of feed with their claws and mouth

parts, disseminating viral particles all over the surrounding water. It is worth noting that the

nephropores are very closely located to the mouth. Besides, when WSSV-containing feed

enters the mouth, it is possible that the feed undergoes a serial passaging of digestive steps,

during which WSSV is liberated from the fed material. Consequently, individual WSSV-virions

could leave the shrimp via the anus or from the degrading bolus, ready to infect shrimp via

the nephropore (Fig. 1).


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Fig. 1. Hypothetical model of shrimp WSSV infection via the nephropore

This thesis did not disprove the per os route as a potential entry point, it did however prove

that it is very inefficient, especially when compared to the nephrocomplex route. It is possible

that under normal conditions the per os route is used by the virus to subclinically infect shrimp

and to keep WSSV present within a population until specific conditions leading to more

frequent urination and to more overt infections. Also, the development of symptoms is most

likely correlated to the WSSV titre that meets susceptible pathways or routes.

7.4. THE SECONDARY DEFENCES OF THE SHRIMPS NEPHROCOMPLEX

When the nephrocomplex is considered as a major entry portal for several pathogens, what

kind of defence can the shrimp mount against invading treats? Of course, the first defence

barrier is the nephropore. With its hard outer cuticular check-valve like structure, the

nephropore remains a very sturdy guardian of the nephrocomplex access. Only when

urinating, this mechanism is briefly compromised. Under normal conditions urination should

not be a big concern. But when a high viral load and/or highly virulent pathogen is present in


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the vicinity of the nephropore in combination with risk factors, the nephropore defence is not

enough on its own. What happens when this first barrier is breached? There is no inner

cuticular lining to prevent pathogens from entering. Fixed phagocytes (podocytes) are present

in the epithelium of the labyrinth and coelomosac but are not yet observed in the rest of the

organ (Doughtie and Rao, 1984; Johnson, 1987; Ueno and Inoue, 1996). Functional

connections have been reported between the nephrocomplex and the lymphoid organ. Inside

the lymphoid organ fixed phagocytes are present, capable of phagocyting pathogens

(Duangsuwan et al., 2008; Rusaini and Owens, 2010) Also, the lymphoid organ expresses

several immune molecules like cathepsin and thrombospondin which would be able to attack

pathogens like Vibrio sp. and viruses (Pongsomboon et al., 2008). In analogy to the lymphoid

organ, the ramifications with no apparent functions could potentially make connections to

the hematopoietic tissue and the latter could thus provide hemocytes to the nephrocomplex’

lumen. Association between the two was observed by proximity, but not by direct connection.

Perhaps other, more gentle, histological techniques could provide more insight. Alternatively,

diapeding hemocytes could enter the nephrocomplex lumen to battle infections. Another

possibility could be that some cells of the nephropore produce or at least allow immune

molecules to pass inside the nephrocomplex lumen. Examples of such proteins are pathogen

recognition receptors containing molecules like lipopolysaccharide and/or β-1,3-glucan

binding proteins (Söderhäll et al., 1990; Lee et al., 2000), lectines (Sharon and Lis, 1989), Toll-

receptors (Arts et al., 2007), serine-protease homologues (Liu et al., 2007), scavenger

receptors (Mekata et al., 2010), DNA- and RNA-binding proteins (Wang et al., 2009), and thio-

ester proteins (Gollas-Galván et al., 2003). The same goes with other antimicrobial proteins

such as penaeidins (Destoumieux et al., 1997), lysozyme (Mai and Wang, 2010), crustins

(Smith et al., 2008), histones (Patat et al., 2004), antilipopolysaccharids (Somboonwiwat et

al., 2005; Liu et al., 2006), and stylicin (Rolland et al., 2010). However, these proteins require

the presence of hemocytes as they depend on these cells to produce them. Another potential

candidate could be hemocyanin which is produced by the hepatopancreas, another organ in

close proximity to the nephrocomplex (Spindler et al., 1992). Finally, a prime candidate is the

clotting protein. It is reported to be expressed by all tissues with exception to hemocytes and

is considered as a first reaction protein against all sorts of pathogens (Yeh et al., 1999;

Maningas et al., 2008). Specific screening of shrimp urine for these molecules should provide

an answer to this question.


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7.5. IMPLICATIONS FOR SCIENCE AND THE INDUSTRY

Regarding management, these proceedings in WSSV-pathogenesis provide ample

opportunities to directly act on managemental level concerning biosecurity. Firstly,

monitoring salinity levels and maintaining those levels should be, if not already, a major

priority for shrimp farmers. Keeping the salinity up to standards and avoiding brisk

fluctuations can be achieved by frequent automated measurements of pond salinity and

avoiding the addition of large volumes of fresh water to the pond (direct rainfall, indirect

rainfall, cleaning water, refreshing the pond water, etc.). Local farmers, operating outdoors

will face severe difficulties in managing the water inflow. A shift to more intensive,

professional farms operating indoors will bring big improvements regarding this fact. Next,

avoiding aggression and the need to establish social dominance, could be profitable. Besides

the other obvious effects of stress, feeding at multiple places in the pond as opposed to one

central point, will reduce fighting and dominant behaviour during the feeding process and

thus less opportunities for WSSV-entry. However, hardly practical in intensive shrimp farming,

one must consider the benefits of adding some kind of sand-like soil or other obstacles for

shrimp to hide and escape aggression or other social events, especially during and after the

moulting process. Nevertheless, such sand-like soil must be bacteriologically justified and thus

it should be possible to clean or regularly replace this soil. Finally, considerations should be

made to artificially select breeding shrimp to create new shrimp generations with the most

growth per moulting cycle, so fewer moulting cycles are required to obtain a similar

harvesting weight.

For several years, researchers try to come up with antiviral therapeutics for white spot

disease. There have been experiments performed, testing the capability of several antiviral

substances (like viral proteins, phytochemicals, antibodies, or RNA interference) with variable

success (Witteveldt et al., 2004; Peraza-Gómez et al., 2009; Nguyen et al., 2018). One

common disadvantage of these methods is that no efficient delivery route exists to get the

anti-viral compound inside the shrimp. These compounds are typically tested using individual

injection of animals, a practice that is hardly convenient in (intensive) shrimp farming. This

thesis can provide a solution for this problem. Experiments can now be set up to investigate

whether antivirals can be delivered when added to the water in hyposaline conditions to allow

these chemicals to enter via the nephropore inside the shrimp to exert its action. An


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additional benefit of this methodology would be that in case of RNA interference and

antivirals with direct mode of action on the virus, its presence in the nephrocomplex can

establish a local protective immunity. Another disadvantage of these antivirals is

sustainability of action. Again, when first applied via salinity shock, bacteria producing

antivirals could potentially be able to settle among the other members of the natural

microflora of the nephrocomplex. Further research hereon is clearly desired.

Previous pathogenesis research was traditionally performed on methodologies following the

injection or per os inoculation. Findings based on these entry routes may be confounded and

thus be done anew with the intra bladder inoculation technology. In fact, time course studies,

comparing the different aspects of WSSV-infection should now be performed with more

detail, comparing the different inoculation routes. Also, other studies with infection aspects

could be revised using intra bladder inoculation.

7.6. CONCLUSIONS

The shrimp nephrocomplex, previously known as the antennal gland, is a very complex organ,

with potentially more functions than previously thought. Its anatomy comprises a vast

network of bladders and tubuli, and spreads from the cephalon to the caudal part of the

thorax and is thus in close contact with the majority of other organs. Whether this has

functional implications, remains to be seen (e.g., lymphoid organ, hepatopancreas, etc.).

Furthermore, it was evidenced that the bladders of the nephrocomplex play a mechanical role

in the moulting and growing process. More research, eluding the full role in these processes

is warranted. Despite the decennia long misconception that the gut is the main entry point

of WSSV, this thesis demonstrated that the nephrocomplex carries this role with much more

efficiency. Risk factors such as salinity shock, stocking density and moulting pose risks for

WSSV-entry. These risk factors all lead to a higher urination frequency and thus to more

opportunities for WSSV-entry via the nephropore into the nephrocomplex where primary

replication transpires.


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7.7. REFERENCES

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Summary

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Summary

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The white spot syndrome virus (WSSV) is a socioeconomic devastating pathogen terrorising

shrimp industry. Beside biosecurity and management measures, no effective measures exist

in the ordeal against WSSV, curative nor prophylactic. The main hindrance hampering

development of such adequate means is a lack of knowledge about pathogen entry and the

primary replication sites. The knowledge on where and how the pathogen enters the shrimp’s

body is crucial information in the struggle to find new ways to combat infectious diseases in

shrimp farming, as well as making advances in WSSV epidemiology. Furthermore, knowing

where and how pathogens enter shrimp provides hints to possible risk factors, while

management strategies can be undertaken to cut the chain of transmission (e.g., animal

isolation, feed supply, waste management, hygiene, vector control, etc.). Finally, when the

method of inoculation mimics the actual, natural way of infection, further pathogenesis

studies can be performed with higher accuracy. Nevertheless, this primary part of

pathogenesis research is too often neglected. Decennia long, researchers assumed that WSSV

entered the host organisms via the per oral route, albeit very inefficiently. Moreover, the gut

is protected by an impenetrable layer of chitin, as is the most part of the shrimp and the lining

epithelium is completely refractive to WSSV-entry. This thesis describes the process of finding

an efficient portal of entry, a corresponding site of primary replication, and the conditions

under which infection is facilitated via that pathway.

In Chapter 1, a comprehensive overview of current relevant knowledge concerning WSSV is

given. This includes an outline of the global significance of WSSV, taxonomy, viral morphology,

and host range and vectors. Most importantly, the current status of knowledge on the

pathogenesis is given. The known transmission mechanisms, pathogen entry, and first barrier

mechanisms protecting shrimp against pathogen entry are highlighted. It is also explained

why the antennal gland, the shrimp’s excretory organ, is considered a prime candidate as

entry portal of WSSV. Finally, a summary of current advances in the knowledge of crustacean

antennal glands is given with the main focus on the penaeid situation. Chapter 2 summarises

the main objectives and methods of the different chapters and the thesis as a whole.

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Currently, the shrimp’s antennal glands’ morphology and anatomy is poorly described.

Therefore, in Chapter 3, a 3D-reconstruction based on histological techniques was applied to

obtain a complete image of the shrimp’s excretory organ. Besides the already known

structures, a rostral continuation of the labyrinth was observed along with a complex of

bladder-like structures caudally of the urinary bladder, reaching as far as the middle of the

hepatopancreas. Regarding the thesis’ objectives, no inner cuticular or other chitinous

protective layer was found, the structure’s walls were found to be easily penetrable, and the

organ was observed to be in close proximity of all other WSSV-susceptible organs and tissues.

Additionally, conclusions and hypothesises considering the basic shrimp biology could be

formulated, e.g., moulting and the urination process. The cellular morphology and the

association with several muscles indicated that the bladders of the nephrocomplex have a

primarily dynamic function. Based on all observations, a new name for the antennal gland

was proposed i.e., “the nephrocomplex”.

In both chapter 1 and chapter 3, links between WSSV-infection, the nephrocomplex, and the

moulting process were unearthed. To further investigate the relationship between the

nephrocomplex (antennal gland) and the moulting process, micro–Magnetic Resonance

Imaging was applied in Chapter 4. Firstly, applying this technique allowed for an in vivo

confirmation of the nephrocomplex constellation as first described ex vivo in chapter 3.

Furthermore, the volume of the bladder structures appeared to be dynamically changing

according to progressing stages of the moulting cycle. Especially around ecdysis, this volume

is at its peak only to gradually decrease to reach a minimum around stage D1. Interesting

conclusions regarding moulting and growing could be drawn. The volume of the caudal

extensions of the lateral compartment of the nephrocomplex could aid the shrimp to break

out of its old shell during ecdysis. Moreover, the volume in the bladders help the shrimp

expand in volume while the new cuticle hardens. Next, the volume inside the bladder will

steadily be replaced by newly formed tissue and organs during post-moulting stages.

Chapter 5 focused on the function of the nephropore to guard the entrance to the

nephrocomplex and to protect against invading pathogenic particles. In other words: how

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effective is the entrance to the nephrocomplex sealed and how can it be penetrated?

Scanning electron microscopy images resulted in the identification of the nephrocomplex

construction as a check-valve like mechanism. Biodynamic ex vivo models were set up and

determined that efflux of fluid (i.e., opening of the nephropore during the mictio) is possible

after a passive build up in pressure to 2 bar, but that the inverse, namely influx of fluid is not

possible even after high manual pressure is exerted. The most important conclusion of this

chapter is that after biodynamic ex vivo simulation of the mictio process, particle influx was

possible. Altogether, this means that the nephropore functions as quite an effective first

defence mechanism against influx of pathogens and is only (briefly) breached during the

process of urinating.

Preceding chapters demonstrated that (i) the nephrocomplex is anatomically and

morphologically suitable as a primary replication site, (ii) the organ is engaged during several

WSSV-infection linked conditions such as moulting and osmostress, and (iii) that urination

poses risk for particles to enter the nephrocomplex. The missing link in the pathogenesis story

was now the actual capacity of pathogens to invade, infect and use the organ as primary

replication site. Chapter 6 deals with artificial and natural infection of shrimp with WSSV and

Vibrio via the nephrocomplex. First, the nephrocomplex was artificially infected using

intrabladder inoculation, which bypasses the check-valve like defence mounted by the

nephropore and was compared to intramuscular injection and per oral inoculation.

Intrabladder inoculation turned out to be over 28 million times more efficient for WSSV in

infecting shrimp as compared to per oral inoculation and only 56 times less efficient when

compared to intramuscular injection, which bypasses all barriers. To demonstrate the broad

range of pathogens to infect shrimp via the nephrocomplex, the experiment was repeated

with the bacterium Vibrio campbellii. Intrabladder inoculation was one billion times more

efficient than peroral inoculation, demonstrating the broad range of pathogens able to cause

disease when entering the nephrocomplex. Next, WSSV-immersion during salinity shock was

employed to simulate a natural infection, i.e., the passing of WSSV through the nephropore

without artificial intervention. Quantitative PCR of urine and hemolymph samples taken at

different time points, showed that urine became positive before the hemolymph did. This is

compelling evidence that WSSV first reaches the urine (and thus the nephrocomplex) before

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the virus can reach the hemolymph and infect all other relevant organs. Additionally, indirect

immunofluorescence staining, confirmed primary replication of WSSV in the ventral urinary

bladder of the shrimp nephrocomplex.

Finally, in the discussion (Chapter 7) all previous chapters were brought together by

formulating an overarching narrative and situating the newly found data, drawn conclusions,

and formed hypothesises within the bigger story of WSSV-entry via the nephrocomplex. Also,

the conditions leading to increased frequencies of mictio were discussed. The latter was

identified as an ultimate risk factor for WSSV-infection. Conditions during which the mictio

frequency is higher are present during post-moulting, salinity shock, episodes of acute stress,

and social behaviour. Based on all data presented in this thesis, a new model for WSSV-

invasion was proposed. The implications of these findings for the industry as well as assists to

future measures and research were provided.

Samenvatting

G.M.A. De Gryse



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185

Het White spot syndrome virus (WSSV) is een socio-economisch enorm relevant pathogeen

dat de garnalen industrie in zijn greep houdt. Behalve bioveiligheid en

managementmaatregelen, bestaan er geen effectieve middelen tegen WSSV. Noch curatief,

noch profylactisch. De ontwikkeling van dergelijke middelen wordt in de weg gestaan door

het ontbreken van relevante kennis op het vlak van pathogenese en meer bepaald over hoe

het pathogeen de garnaal kan binnendringen en het primaire replicatie weefsels kan

bereiken. De informatie over hoe en waar het virus de garnaal binnenkomt, is cruciaal om

nieuwe bestrijdingsmiddelen te vinden, maar ook om vorderingen te maken op het gebied

van de epidemiologie. Bovendien, levert deze informatie hints over mogelijke risico factoren

voor WSSV-infectie. Managementmaatregelen kunnen dan genomen worden om de

transmissie keten te onderbreken (vb. quarantaine, voeding, mest- en afvalverwerking,

hygiëne, controle van vectoren, etc.). Wanneer de inoculatiemethode de natuurlijke weg van

infectie het beste nabootst, kunnen pathogenese studies met een hogere accuraatheid

verlopen. Niettegenstaande is dit cruciale onderdeel van het pathogenese onderzoek al te

vaak verwaarloosd. Decennialang zijn onderzoekers er van uit gegaan dat WSSV de garnaal

via het maagdarmstelsel (peroraal) infecteert. Echter, deze infectieroute blijkt zeer inefficiënt

te zijn en bovendien is de binnenzijde van het spijsverteringsstelsel volledig beschermd door

een ondoordringbare chitineuze laag en is het epitheel hier refractair voor WSSV-infectie.

Deze thesis beschrijft de zoektocht naar het vinden van een efficiënte intredepoort, een

bijbehorende primaire replicatie site en de omstandigheden waarbij infectie wordt

gefaciliteerd.

In Hoofdstuk 1, wordt er een uitgebreid overzicht gegeven over de relevante kennis

betreffende WSSV. Het gaat hierbij onder andere over de globale spreiding, taxonomie, virale

morfologie, gevoelige gastheren en vectoren. Bovenal wordt de huidige kennis over de

pathogenese beschreven. De focus ligt hierbij op gekende overdrachtsmechanismes,

pathogeen intrede en primaire verdedigingsmechanismes. Ook wordt er uitgelegd waarom

de antennale klier wordt verdacht als intredepoort voor WSSV. Om te eindigen wordt er ook

een overzicht gegeven over de huidige stand van zaken over de antennale klier bij

schaaldieren. Daarbij wordt er speciale aandacht geschonken aan de situatie bij penaeide

garnalen. Hoofdstuk 2 synthetiseert het geheel van hoofddoelstellingen en methoden over

de verschillende hoofdstukken.

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186

Tegenwoordig is de morfologie van de antennale klier bij garnalen slecht beschreven. Daarom

wordt er in Hoofdstuk 3 een 3D-reconstructie gemaakt van dit excretie orgaan die gebaseerd

is op een histologische methodologie. Dit liet ons toe om een totaalbeeld te verkrijgen van de

anatomie en morfologie van de antennale klier. Naast de reeds gekende structuren, werden

er immers een rostrale verderzetting van het labyrint ontdekt, samen met een complex van

blazen en zakken meer naar caudaal. Dit blazencomplex reikte tot aan het midden van de

hepatopancreas. Hierbij werd er bovendien geen cuticulaire of andere chitineuze aflijning

teruggevonden. Ook was de dikte van de cellagen zeer beperkt en blijkt de structuur zich dicht

tegen alle andere WSSV-gevoelige orgaan te situeren. Daarnaast, deed de nieuwgevonden

morfologie andere hypothesen omtrent de basisbiologie van de garnaal bovendrijven zoals

vervelling en verzorging van de kieuwen. De cellulaire morfologie en de associatie met

verschillende spieren, wijzen op een grotendeels dynamisch aspect van het blazencomplex.

Gebaseerd op alle observaties, werd er tenslotte een nieuwe naam voorgesteld voor de

antennale klier: “het nefrocomplex”.

Zowel in hoofdstuk 1 als hoofdstuk 3, werden er linken gelegd tussen enerzijds WSSV-infectie

en het vervellingsproces anderzijds. Om de relatie tussen het vervelingsproces en het

nefrocomplex (antennale klier) verder de onderzoeken, werd er micro magnetische

resonantie beeldvorming aangewend in Hoofdstuk 4. Vooreerst leverde het toepassen van

deze methodologie een in vivo bevestiging op van het 3D-model uit hoofdstuk 3. Verder, bleek

ook het volume van het blazencomplex te variëren naargelang de verschillende

vervellingsstadia. Vooral rond het eigenlijke vervellingsmoment bleek de vulling op het

maximum te zijn. Na de vervelling gaat dit gradueel verminderen om een minimum te

bereiken rond het D1-stadium. Interessante conclusies konden worden getrokken

betreffende het vervellings- en groeiproces. Het toegenomen volume in de caudale extensies

van het laterale compartiment van het nefrocomplex kan de garnaal mechanisch helpen om

uit zijn oude schaal te breken. Bovendien zal het volume de garnaal helpen om te expanderen

terwijl de nieuwe schaal nog zacht is. Finaal zal de inhoud van het blazencomplex vervangen

worden door het volume dat nieuwgevormd weefsel zal innemen.

Hoofdstuk 5 focust op de capaciteit van de nefropoor om de toegang tot het nefrocomplex

te bewaken en te beschermen tegen invaserende partikels. M.a.w. hoe effectief wordt de

toegang tot het nefrocomplex afgesloten en hoe kan dit systeem worden omzeild? Scanning

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elektronenmicroscopie identificeerde de nefropoor als een soort terugslagklep.

Biodynamische ex vivo modellen werden ontwikkeld en toonden dat efflux (openen van de

nefroporie tijdens de mictio) mogelijk is door een passieve opbouw van druk (2 bar), maar dat

de omgekeerde beweging nl. influx onmogelijk is, zelfs met een zeer hoge druk. De

belangrijkste conclusie van dit hoofdstuk, is dat wanneer er een mictio wordt gesimuleerd,

dat er wel vloeistof en partikels de garnaal binnen geraken via de nefroporie. Alles in acht

genomen, betekent dit dat de nefroporie een zeer effectieve afsluitklep is, die enkel tijdens

het urineren kan omzeild worden. Condities waarbij garnalen vaker urineren, zijn dus risico

factoren op WSSV-uitbraken.

In vorige hoofdstukken werd duidelijk dat (i) het nefrocomplex anatomisch en morfologisch

geschikt is als een primaire replicatiesite, (ii) het orgaan betrokken is tijdens verschillende

aan-WSSV-infectie-gelinkte-condities zoals de vervelling en osmostress en (iii) dat het

urineren een risico inhoud voor partikelinvasie via de nefroporie. Om het pathogenese

verhaal af te ronden is het nodig om effectief een infectie van en via het nefrocomplex aan te

tonen. Hoofdstuk 6 handelt over de artificiële en natuurlijke infectie van WSSV en Vibrio in

garnalen via het nefrocomplex. Eerst werd het nefrocomplex artificieel geïnoculeerd via een

intrablaastoediening van beide pathogenen apart. Deze inoculatiemethode omzeilt het

terugslagklepmechanisme van de nefroporie die de nefrocomplex beschermt tegen

partikelinvasie. De Intrablaasinoculatie werd vergeleken met intramusculaire en perorale

injecties. Hieruit blijkt dat de intrablaasinoculatie meer dan 28 miljoen keer efficiënter is dan

perorale inoculatie en slechts 56 keer minder dan intramusculaire injectie. Het experiment

werd met succes herhaald om de brede range aan pathogenen aan te tonen waaraan het

nefrocomplex gevoelig is. Vervolgens werd osmosestress als risicofactor misbruikt om een

natuurlijke infectie teweeg te brengen. Daarbij werd om de twaalf uur urine- en

hemolymfestalen genomen. Een kwantitatieve PCR-assay bracht aan het licht dat de urine

eerst positief werd vooraleer WSSV in de hemolymfe kon worden teruggevonden. Dit is een

sterk bewijs dat WSSV eerst via de nefroporie het nefrocomplex bereikt, dat er daar een

primaire replicatie plaatsvindt om vervolgens de hemolymfe te bereiken alwaar een

algemene infectie opgang wordt getrapt. Ter bevestiging, werd er met indirecte

immunofluorescentie een primaire infectie van het nefrocomplexepitheel aangetoond.

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Alle voorgaande hoofdstukken werden in Hoofdstuk 7 samengebracht in een algemene

discussie. De nieuwgevonden data, getrokken conclusies en gevormde hypotheses worden

geduid binnen een overkoepelend verhaal over de pathogenese en invasiemechanismen van

WSSV via het nefrocomplex. De verschillende condities die leiden tot het verhogen van de

mictiofrequentie werden ook bediscussieerd. Dat laatste werd in deze thesis immers in

verband gebracht met een verhoogde kans op WSSV-infectie. Voorbeelden van zo’n condities

of risicofactoren zijn: vervelling, acute stress, osmostress en sociaal gedrag. Gebaseerd op alle

data gepresenteerd in deze thesis werd er tenslotte een nieuw model voorgesteld voor WSSV-

invasie. Deze bevindingen kunnen de industrie op weg helpen naar minder (WSSV)

besmettingen door enkele concrete managementmaatregelen. Ook werd de voorzet gegeven

tot verdere potentiële onderzoeken en methodes om de garnalen industrie te beschermen

tegen pathogenen. Gebaseerd op alle data die in deze thesis werd beschreven, werd een

nieuw model voor WSSV-invasie voorgesteld.

Curriculum Vitae and publications

G.M.A. De Gryse




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Gaëtan De Gryse werd op 30 oktober 1990 geboren in Oostende. In 2008 behaalde hij het

diploma hoger secundair onderwijs in de richting Wetenschappen-Wiskunde aan het Sint-

Andreasinstituut te Oostende. Tijdens het laatste jaar van zijn middelbare studies werd hij

laureaat van de Vlaamse Biologie Olympiade. Vervolgens werden hogere studies aangevat

aan de Universiteit Gent waar hij in 2015 met onderscheiding afstudeerde als master in de

Diergeneeskunde optie onderzoek, cluster grote huisdieren. Voor zijn masterthesis ontving

hij de Thesis Award Marine Sciences uitgereikt door het Vlaams Instituut voor de Zee. Na zijn

studies diende Gaëtan een projectaanvraag in bij het toenmalige Instituut voor Wetenschap

en Technologie (IWT) en verwierf daarop een doctoraatsbeurs ‘strategisch basisonderzoek’

van het Fonds Wetenschappelijk Onderzoek (FWO). Onder begeleiding van Prof. Hans

Nauwynck en Prof. Peter Bossier startte hij aldus een doctoraatsonderzoek aan waarbij de

pathogenese van het WSSV werd bestudeerd. Het relaas daarvan is terug te vinden in deze

thesis. Hij is auteur en coauteur van publicaties in verschillende internationale tijdschriften

en schreef twee hoofdstukken in het internationaal gewaardeerde en druk geraadpleegde

garnalenhandboek die hij in het begin van zijn doctoraatstudie zelf frequenteerde voor

informatie rond de wetenschap van de garnalenkweek. Hij begeleidde drie honours-

studenten en twee masterproefstudenten.

Gaëtan De Gryse was born on the 30th October 1990 in Ostend, Belgium. In 2008, he obtained

his high school degree, majoring science, and math at the Sint-Andreas institute also in

Ostend. During his senior year, he was laureated at the Flemish Biology Olympiad. Next, he

commenced his higher education at the University of Ghent. In 2015, he graduated with

distinction as Master of Veterinary Medicine, option research, cluster large animals. For his

master’s thesis, Gaëtan received the VLIZ Thesis Award for Marine Sciences from the Flanders

Marine Institute. Next, he submitted a project application with the former Institute for

Science and Technology (IWT) and received funding for his PhD-research at the Laboratory of

Virology at the Faculty of Veterinary Medicine from the Research Foundation - Flanders

(FWO). Under the tutelage of Prof. Hans Nauwynck and Prof. Peter Bossier, he commenced


194

researching the WSSV pathogenesis in shrimp. Expeditions to several other scientific

disciplines was needed to achieve his goals. Furthermore, he authored and co-authored in

international journals, as well as authored two chapters in the leading Shrimp Book, which he

frequently consulted during the first years of his doctoral research. He promoted 3 honours

students and 2 master thesis students.


195

SCIENTIFIC PUBLICATIONS (5) 2020 The shrimp nephrocomplex serves as a major portal of pathogen entry and

is involved in the molting process PNAS; Proceedings of the National Academy of Sciences of the United States of America (2020) Gaëtan M. A. De Gryse, Thuong Van Khuong, Benedicte Descamps, Wim Van Den Broeck, Christian Vanhove, Pieter Cornillie, Patrick Sorgeloos, Peter Bossier, and Hans J. Nauwynck https://doi.org/10.1073/pnas.2013518117

2018 Establishment of porcine enterocyte/myofibroblast co-cultures for the growth of porcine rota- and coronaviruses Scientific Reports (2018) Tingting Cui, Sebastiaan Theuns, Lowiese Desmarets, Jiexiong Xie, Gaëtan M.A. De Gryse, Bo Yang, Wim Van Den Broeck, and Hans J. Nauwynck https://doi.org/10.1038/s41598-018-33305-1

2016 Upregulation of endothelial cell adhesion molecules characterizes veins

close to granulomatous infiltrates in the renal cortex of cats with feline infectious peritonitis and is indirectly triggered by feline infectious peritonitis virus-infected monocytes in vitro Journal of General Virology (2016) Delphine D. Acar, Dominique A. J. Olyslaegers, Annelike Dedeurwaerder, Inge D. M. Roukaerts, Wendy Baetens, Sebastiaan Van Bockstael, Gaëtan M. A. De Gryse, Lowiese M. B. Desmarets, and Hans J. Nauwynck https://doi.org/10.1099/jgv.0.000585

2016 Kinetic analysis of internalization of white spot syndrome virus (WSSV) by hemocyte subpopulations of penaeid shrimp, Litopenaeus vannamei (Boone), and the outcome for virus and cell Journal of Fish Diseases (2016) Vo Van Tuan, Gaëtan M.A. De Gryse, Khuong van Thuong, Peter Bossier, Hans J. Nauwynck https://doi.org/10.1111/jfd.12482

2015 Virus replication cycle of White spot syndrome virus in secondary cell cultures from the lymphoid organ of Litopenaeus vannamei Journal of General Virology (2015) Wenfeng Li, Lowiese M. B. Desmarets, Gaëtan M. A. De Gryse, Sebastiaan Theuns, Vo Van Tuan, Khuong Van Thuong, Peter Bossier, and Hans J. Nauwynck https://doi.org/10.1099/vir.0.000217


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BOOK CHAPTERS (2)

2021 On shrimp pathogen entry: the case of WSSV as a guide to other shrimp

pathogens The Shrimp book volume 2 (in print) Gaëtan M.A. De Gryse, Mathias Corteel, Cesar Escobedo-Bonilla, Thuong Van Kuong, Peter Bossier, Patrick Sorgeloos, and Hans J. Nauwynck

2021 Functional morphology of the nephrocomplex

The Shrimp book volume 2 (in print) Gaëtan M.A. De Gryse and Hans J. Nauwynck

Acknowledgements-Dankwoord

Acknowledgement -Dankwoord

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Completing a doctorate is a feat no one ever accomplishes alone. Therefore, I would like to

dedicate this little part of the thesis to everybody who once helped me to get me to this place.

Whilst writing this acknowledgement, an uncomfortable feeling keeps creeping up to me that

I just might forget some of you or even that my words do not remotely match the merits of

everybody involved. For the unacknowledged soldiers: thanks!

Geen opportuniteit, geen doctoraat. Prof. Nauwynck, Hans voor de ingewijden, zag na mijn

masterthesis aan zijn labo voldoende potentieel in mij om mij destijds een kans te geven om

samen een projectaanvraag voor te bereiden, in te dienen en te verdedigen. Als we één iets

gemeenschappelijk hebben, dan is het wel onze voorliefde voor de meeste gekke hypothesen

en ditto experimenten. Daarin vond ik de vrijheid die mij motiveerde. Bedankt ook om het

arty farty (“but what does it mean”) van de hedendaagse wetenschap in perspectief te

plaatsen met een vleugje gezond verstand. Uitgaan van eigen kracht, en dan vooral van de

kracht van de veterinairs.

Waar Hans mijn wilde kant aanstookte, was er toch steeds de rustige, realistische

temporisering van Peter. Ik herinner mij nog de vele (slapeloze) nachtjes op een oude

luchtmatras in het oude gebouw van het ARC waar ik garnalen onderdompelde in fluogroen

water tjokvol genetisch gemanipuleerde vibrio’s. Peter, jouw immense kennis van de

aquacultuur, garnalen en bacteriën, stuurden mijn experimenten meer dan eens van wild

cowboy verhaal naar realistische en haalbare opzetten.

Nico, door omstandigheden hebben wij niet zo veel kunnen samenwerken als ik had gehoopt

bij het begin van dit doctoraat. Maar jouw rol was toch cruciaal in de wijze en broodnodige

woorden die jij mij schonk.

A big thanks as well to the members of the examination committee as their comments

ultimately improved the quality of this thesis and urged me to think on a few extra

conclusions.

Zeer veel dank ben ik verschuldigd aan mijn virologische pleegouders Bas, Lowiese, Delphine,

Inge en Isaura die mij destijds adopteerden uit de krochten van de kelders onder de

hoogbouw en mij doopten tot “king under the mountain”. Ze namen mij mee naar de warmte

van de virologische enclave op het parasitaire verdiep.


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Bas, wat jij voor mij betekend hebt, kan ik met geen woorden vatten. Van toen ik nog master

student was, was jij het die mij onder jouw vleugels hebt genomen en ingewijd in de geheimen

van de (q)PCR en wat nog allemaal. Jij hebt toen nog tijdens het afwerken van jouw eigen

doctoraat de tijd genomen om tijdens de communie van jouw familielid mijn discussie na te

lezen. Die toewijding zal ik nooit vergeten. Het samen met jouw rondtjoolen doorheen het

laboratorium was een plezier die ik niet zal vergeten. We zien elkaar zeker nog op en naast

de fiets/minion.

Lowiese, mijn overbuur. Wat voor Bas geldt, gaat zeker ook op voor jou. In het begin van mijn

doctoraat kon ik steeds bij jou met al mijn gekke hypothesen en rare vragen. Ik betrap mij er

vandaag de dag nog op dat ik binnensmonds jouw liedjes zit te zingen. Dikke duim! Het gat

dat je hebt nagelaten toen je naar Frankrijk vertrok is nog steeds niet opgevuld binnen de

DI04. Onze trip naar het REGA-instituut zal ik ook niet gauw vergeten: Peppi en Kokki in

Leuven!

Ook aan jou Delphine, heb ik veel te danken, je maakte mij wegwijs in het labo en ik had

steeds jouw aandacht als mijn concentratie kelderde. We hebben elkaar gesteund als het

moeilijker ging, maar ook veel plezier beleefd! Als je ooit nog een lucky charm nodig hebt bij

het electroporeren van jouw E. coli’s, roep maar! Wat wij allemaal hebben uitgestoken: te

veel om te herinneren.

Isaura, onze kritische rots in de branding. Meer dan eens zette jij mij met de voeten op de

grond nadat je mij roosterde over mijn proefopzet. Daar heb ik toen, en nu nog steeds, veel

aan gehad. Inge, zonder veel te zeggen droegen jouw woorden wellicht het meeste gewicht.

In dat warme nest vond ik ook mijn partner in crime Sebastiaan ‘Ryan Gosling’ Bockstael. Ik

vond het fijn om naast jou te werken en samen het labo te ontdekken. Wat hebben wij

gelachen dag na dag. En gegaan met die banaan… Ongelofelijk. Samen met Delphine en Louis

startten we onze eigen KFFC-quizploeg waarvoor menig anderen trilden in hun schoenen. Als

die Corona-toestanden wat gaan liggen zijn, zullen we zeker heropstarten!

Ytse, het zooitje ongeregelde waarover ik net heb geschreven (en waar ik zeker bij hoor)

moest meer dan eens in het gelid gehouden worden. Ik ben zeker dat jij onder jouw haar een

punthoofd hebt verborgen waarvan ik de schuldige ben. Mijn antwoord was meer dan eens:

“met grote creativiteit, komt grote chaos”… niet dat het ooit pakte maar soit. Het was met


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veel spijt dat ik je zag vertrekken naar de humane virologen, ze weten nog niet half hoe veel

geluk ze daar hebben Bedankt voor al het secure qPCR-werk! Op jou kan je rekenen.

Ook Wendy, Veerle, Kasper, Eleni, Jonathan, Marthe en Robin bedankt voor alle momenten

die we samen hebben gekend. Also, thanks to Dayoung, with whom I had the pleasure of

having adjoining desks. I will always keep you in fond memory. And I will never forget how

you have introduced me to the confocal microscope. I wish you a lot of success in your future

career. Tingting, Jason, Vishi, Bo, Charlie, and Kevin, it was a pleasure to work alongside you

in the lab and I hope one day, I could come and visit you again. Liping and Mostafa, I have

only had a short time to work with you, but I hope you will continue the work I started. I wish

you all the best in the future and if you would need any advice, I will be happy to help you

where I can.

To Noémie, Eline, Nick and all other current phd students of the lab: I wish you a lot of courage

and luck in finishing your PhD.

Tijdens mijn doctoraat heb ik vele laboratoria rondgezworven waar ik steeds mensen

terugvond die mij hielpen en waarop ik kon bouwen: prof. Simoens, Prof Wim Van Den

Broeck, Prof. Pieter Cornillie, bedankt voor de hulp met de histologische,

elektronenmicroscopische en morfologische hulp. Prof. Annemie Decostere en Prof. Koen

Chiers, bedankt voor de hulp bij het maken van de garnalencoupes. Prof. Katrien

Vanderperren, heel veel dank voor het constructieve RX-onderzoek. Ook de vrienden van het

Infinity lab mogen niet onvermeld blijven. Chris en Benedicte, ik herinner mij nog steeds jullie

gezichten toen Hans en ik voor het eerst bij jullie binnen stuikten met ons emmertje garnalen.

Jullie waren van het eerste moment enthousiast en mee in het verhaal. Ook de mensen van

het ARC, Tom, Jorg, Matthieu, Margriet en Gilbert verdienen een bedankje voor de vele

logistieke en administratieve steun. Ook Tim van CMET, bedankt voor de hulp bij de

genoomanalyses.

Aux nouveaux collègues du service ENZOVEB de Sciensano, merci de m’avoir accueilli. Il me

semble que ce soit un très grand plaisir et opportunité de travailler avec vous et de faire des

projets ensemble. A partir de maintenant, je pourrai passer plus de temps avec vous. Brigitte,

ik geloof echt dat ik in een mooie omgeving ben geland. Bedankt om mij de tijd en ruimte te

laten om deze thesis af te werken. Ik sta te popelen om met jullie hoge toppen te scheren.


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Beste vrienden en familie, jullie zijn te talrijk om individueel te lauweren (tijdsnood, weet je

wel ). Oostende, Beveren, Diergeneeskunde, Buren, judo en judo. “Een mens is tot op

zekere hoogte het product van zijn omgeving”. Hippocrates wist het al, en ik zag het

bevestigd. Door heen heel mijn leven ben ik voortdurend warme en interessante mensen

tegengekomen. Sommigen kom ik nog steeds tegen, anderen nu wat minder. We kunnen

voortdurend leren van elkaar en ik hoop dit in de toekomst verder te kunnen zetten. Niets

beters om de gedachten te verzetten in de compagnie van goede vrienden, met of zonder

eten, drank, feest, skilatten, lasagnes, wurgingen of “karrekes”.

Véronique en Erik, zeer bedankt voor de steun de afgelopen jaren en de tweede thuisbasis in

Vrasene City. Ik ben jullie erkentelijk voor de hartelijke opname in het gezin. Niet te vergeten:

bedankt voor de inwijding in de kunst van het fietsen en de mechaniek van het onderhoud.

Mama en papa, doorheen de jaren heb ik jullie ongetwijfeld vele grijze haren bezorgd,

misschien zijn er her en der wel een paar extra uitgevallen door mijn toedoen. Bedankt om

mij te blijven steunen, mij alle kansen te geven die ik nodig had en om in mij te geloven.

Zonder jullie weet ik niet waar ik zou staan, maar het was zeker niet hier.

Ameline, ik was je niet vergeten want jij bent het begin en het einde. Bedankt voor alles wat

je al voor mij deed en ook bedankt voor alles wat je nog zal doen. We vormen nu al een tijdje

een tandem, en hoe verder wij fietsen, hoe beter wij elkaar aanvullen. Synergisme van de

bovenste plank. Nu dit doctoraat afgerond is, zijn we eindelijk klaar om de volgende stappen

in ons leven te nemen. Graaf al maar diep in uw portemonnee en ovaria, dan kunnen we er

snel aan beginnen!

Bedankt, Thanks, Merci,

Gaëtan,

Merelbeke 25/10/2021

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