eukaryotic pathogens (chytridiomycota and oomycota ...

EUKARYOTIC PATHOGENS (CHYTRIDIOMYCOTA AND OOMYCOTA) INFECTING MARINEMICROPHYTOBENTHIC DIATOMS – A METHODOLOGICAL COMPARISON1

Bettina Scholz2,3,4

Institute of Chemistry and Biology of the Marine Environment, University of Oldenburg, Schleusenstrasse 1, Wilhelmshaven

26382, Germany

Frithjof C. K€upper

Oceanlab, University of Aberdeen, Main Street, Newburgh AB41 6AA, UK

Wim Vyverman

Department of Biology, Section of Protistology and Aquatic Ecology, University of Ghent, Krijgslaan 281 S8, Ghent 9000, Belgium

and Ulf Karsten

Institute of Biological Sciences, Applied Ecology & Phycology, University of Rostock, Albert-Einstein-Strasse 3, Rostock 18059,

Germany

Using sediment samples from the Solth€orn tidalflat (southern North Sea, Germany), collected inbi-weekly intervals from June to July 2012, a rangeof qualitative and quantitative screening methodsfor oomycete and chytrid pathogens infectingbenthic diatoms were evaluated. Pre-treatment ofsediment samples using short ultrasound pulses andgradient centrifugation, in combination withCalcoFluor White, showed the best results in thevisualization of both pathogen groups. The highestnumber of infected benthic diatoms was observedin mid July (5.8% of the total benthic diatomcommunity). Most infections were caused bychytrids and, in a few cases, oomycetes (LagenismaDrebes (host: Coscinodiscus radiatus Ehrenberg) andEctrogella Zopf (hosts: Dimeregramma minor inPritchard and Gyrosigma peisonis). Among thechytrids, sporangium morphology indicated thepresence of five different morphotypes, infectingmainly epipelic taxa of the orders Naviculales (e.g.,Navicula digitoradiata) and Achnanthales (e.g.,Achnanthes brevipes Agardh). The presence ofmultiple pathogens in several epipelic diatom taxasuggests a significant role for fungal parasitism inaffecting microphytobenthic diatom succession.

Key index words: benthic diatoms; CalcoFluor White;chytrids; fluorescein isothiocyanate-labeled wheat-germagglutinin; oomycetes; sporangia; stainingmethods

Abbreviations: AF, acid fuchsin; CR, Congo red;CW, CalcoFluor White; FITC, fluorescein isothiocya-nate; LCB, Lactophenol-cotton blue; MMS, Mayer’sMucicarmine stain; MPB, microphytobenthos; NAG,N-acetylglucosamine; NR, neutral red; PI, propidi-um iodide; TB, Trypan Blue; WGA, wheat-germ agg-lutin

Intertidal and shallow subtidal sediments are char-acterized by dense populations of benthic microal-gae, the so-called microphytobenthos (MPB;Paterson and Hagerthey 2001). These organismsplay a key role in coastal ecosystem functioning, andcontribute significantly to the primary productionin littoral zones (Pinckney and Zingmark 1993).They are a major food source for the zoobenthos,and even for commercially important fish and shell-fish stocks as well as for migratory bird populations(e.g., Hillebrand et al. 2002). Most intertidal flatsare mainly colonized by diatom-dominated biofilms(MacIntyre et al. 1996, Middelburg et al. 2000, Pat-erson and Hagerthey 2001), which are usually com-posed of pennate forms, being either epipsammicor epipelic (e.g., Mitbavkar and Anil 2002).Besides abiotic parameters (e.g., Admiraal et al.

1984, Thornton et al. 2002), biotic factors such ascompetition for light and nutrients and predator–prey interactions have been shown to play key rolesin the community structures of benthic diatoms(Thornton et al. 2002). In the marine environment,a growing body of evidence points additionally toparasites as key players in the control of populationdynamics and overall ecosystem structure (Gachonet al. 2010). Several studies on planktonic microalgae(e.g., Chambouvet et al. 2008) and benthic brownmacroalgae (K€upper and M€uller 1999, K€upper et al.

1Received 11 May 2014. Accepted 31 July 2014.2 Present address: BioPol ehf., Einb�uastig 2, Skagastr€ond 545,

Iceland.3Present address: Faculty of Natural Resource Sciences, University

of Akureyri, Borgir v. Nordurslod, Akureyri IS 600, Iceland.4Author for correspondence: e-mail [email protected] Responsibility: P. Kroth (Associate Editor)

J. Phycol. 50, 1009–1019 (2014)© 2014 Phycological Society of AmericaDOI: 10.1111/jpy.12230

1009

2006, Gachon et al. 2009, Strittmatter et al. 2013)have shown the strong impact of eukaryotic patho-gens such as oomycotes and chytridiomycotes ontheir hosts’ ecology (e.g., Bruning 1991, Ibelingset al. 2004, Kagami et al. 2007, Gleason et al. 2008,Sekimoto et al. 2008a,b). In particular, oomycetesinfecting marine phytoplankton comprise severalmarine representatives such as Lagenisma coscinodisciDrebes, which was reported as an endobiotic parasiteof the centric diatom Coscinodiscus centralis Ehren-berg from the North Sea (Drebes 1966, 1968, Gotelli1971). Furthermore, the endoparasitic, saprolegnia-ceous oomycete Ectrogella Zopf is a parasite in dia-toms and, according to Sparrow (1969), outbreaks ofEctrogella perforans Petersen may attain epidemic pro-portions in the marine pennate diatom LicmophoraAgardh. In contrast, chytrid infections seem to bemost common among large species of freshwaterphytoplankton that are fairly resistant to zooplanktongrazing (Sommer 1987). Over 90% of all host cells ina population may be infected, and every infectioncommonly leads to the death of the host cell (Canterand Lund 1951, Ibelings et al. 2004, 2011). From brack-ish and marine ecosystems, only few representatives ofOlpidium (Braun) Rabenhorst andRhizophydium Schenkhave been described as parasites of small marineplanktonic green algae and diatoms (Gleason et al.2011).

The life cycles of oomycetes and chytrids beginwith the attachment of a motile zoospore to thesurface of an algal host cell. Besides differences inzoospore morphology, there are fundamental differ-ences between both groups of parasites. Chytrids aretrue fungi, while oomycetes are stramenopiles, a het-erotrophic sister group, for example, brown algaeand diatoms (Gleason et al. 2011). In consequence,for example, chytrids are characterized by chitina-ceous cell walls, whereas oomycetes have predomi-nantly cellulosic walls (consisting mainly of 1,3-b-glucans, some 1,6-b-glucans and 1,4-b-glucans). Chi-tin, which is a major constituent of fungal cell walls,has been detected in small amounts in only a fewoomycetes (Latijnhouwers et al. 2003). Due theirsmall and holocarpic thalli, microscopic identifica-tion of these parasitic species is not easy and oftenrequires ultrastructural characterization of zoosporesfor absolute certainly (Barr 1981, Longcore 1995,Hanic et al. 2009, Letcher and Powell 2012).

Detection of eukaryotic pathogens infecting phyto-plankton species can be difficult, and different stain-ing methods exist. Among these CalcoFluor White(CW) and wheat-germ agglutin (WGA) conjugatedwith fluorescein isothiocyanate (FITC) followed byobservation unan epifluorescence microscope arefrequently used for the visualization of fungal para-sites (e.g., M€uller and Sengbusch 1983, Rasconiet al. 2009, Marano et al. 2012). For benthic sam-ples, there is the additional challenge to separateintact cells from inorganic and organic particles.Here, the lens-tissue method is routinely used for

collecting epipelic diatoms (e.g., De Jonge 1980),whereas for the sometimes abundant epipsammictaxa, no standard method exists. In this study anarray of staining methods was compared whichallowed the identification of these pathogens usingdifferent enrichment and staining procedures. Spe-cifically, the detection of fungal pathogens in diluteduntreated surface sediment samples was comparedwith those that had been pre-treated with ultrasonicand gradient centrifugation. In addition, infectionrates by eukaryotic parasites of the total microphyto-benthic diatom communities and the impact onindividual taxa were also calculated, depicting forthe first time qualitative and quantitative informa-tion about the occurrence of such pathogens in themarine benthic realm.

MATERIALS AND METHODS

Study site and sample collection. The Solth€orn tidal flat islocated in the eastern part of the Inner Jade, near the villageof Tossens in Lower Saxony, Germany (53°340 2.03″ N; 8°13054.66″ E). One station was sampled at bi-weekly intervals dur-ing low tide from June 16th until July 14th 2012. Triplicatesurface samples of sediment were obtained by inserting8.5 cm diameter plastic petri dishes into the sediment to adepth of 1 cm. All samples were stored at 4°C � 2°C untilfurther processing in the laboratory.

Chemicals. If not otherwise mentioned, all chemicals usedin this study were of the highest purity from Sigma/AldrichChemical Co. (Sigma-Aldrich Laborchemikalien GmbH,Seelze, Germany).

Treatment of sediment samples. The sediment samples wereprepared in four subsequent steps (Fig. 1). In a first step,three sediment subsamples (volume: 1 cm3) were eachdiluted in 40 mL sterilized artificial seawater, using TropicMarin� (GmbH Aquarientechnik, Wartenberg, Germany),with a salinity of 30 and a pH of 8.0, respectively (Fig. 1A).To remove epipsammic diatom taxa from the sediment parti-cles, ultrasonic pulses of 3 9 2 s were used in a second step(ultrasonic processor UP50 Dr. Hielscher GmbH, Tetlow,Germany, amplitude of 40% at 0.5 s intervals, Fig. 1B). Subse-quently, the mixtures were separated in a third step with aslightly modified version of the technique described by DeJonge (1979) (Fig. 1C). The procedure followed is based ondensity gradient centrifugation of the samples in Ludox-TM.Benthic diatom species were harvested from the 70% Ludox-TM layer, washed twice with 2.5 mL of the seawater solutionby centrifugation (10 min at 500 rpm, Centrifuge EBA 20,Hettich GmbH, Tuttlingen, Germany) and placed in glassUterm€ohl counting chambers in a fourth step (100 lL,Fig. 1D). Microscopic observation of the residues (sedimentparticles) using epifluorescence confirmed the almost com-plete release of the cells (Olympus BX51, equipped with aBX-RFA reflected fluorescence system, Hamburg, Germany).800 lL (8 9 100 lL) of the samples from steps one andtwo (after sonification) were used for a first screeningusing autofluorescence and the staining methods describedbelow.

Staining methods. Both parasites attached to diatom cells(epibiotic) as well as endobiotic ones were stained accordingto different methods and protocols (see Table 1). In total tenstaining methods were tested at the beginning of this study,using samples from each step of the sediment preparation(cf. Fig. 1) of which only the FITC-WGA and CW protocolswere modified. For the use of FITC-WGA, 3 mL of borate

1010 BETTINA SCHOLZ ET AL.

buffer (10 mM; pH 7.4) were added to the samples. For thepreparation of a 1.0 mg � mL�1 stock solution, 5.0 mg oflyophilized FITC-WGA in 5.0 mL of phosphate-buffered sal-ine (PBS) were dissolved and stored at �18°C (stable for1 month). For the assay, the stock solution was diluted inPBS to a final concentration of 7.0 lg � mL�1. 1 mL of thissolution was used for labeling. After incubation (30 min;25°C) on a rotary shaker (130 rpm), samples were rinsedtwice with fresh buffer, diluted to a final volume of 5 mL andexamined at 4009 magnification. In contrast, 150 lL of 10%KOH solution and 150 lL of 0.2% CW were added to 1 mLsamples in Uterm€ohl glass counting chambers and diluted toa final volume of 5 mL, after which the samples were incu-bated for 10 min at room temperature. All assays were con-ducted in glass Uterm€ohl counting chambers, because someof the staining agents (e.g., WGA) were observed to bind toplastic material. The samples were counted at 4009 magnifi-cation unan inverted fluorescence microscope with UV excita-tion and also with visible light (Olympus IX53). Diatomspecies were identified as far as possible, enumerated andassigned to taxonomic entities. Abundance was calculated asthe number per cm2 sediment surface. The results of the cellcounts for the June and July samples (June 6, July 1 and 14,2012), comprising all staining methods used, are given in Fig-ure 2.

Identification of eukaryotic pathogens and their hosts. Sub-sam-ples (3 9 50 lL) from all staining preparations weremounted in Slowfade-Light antifade solution (MoBiTec,G€ottingen, Germany), whereas in parallel aliquots(3 9 500 lL and 10 lL, respectively) of the unstained sedi-ment samples after sonification and the enrichment step weredried and embeded in Euparal (Chroma Gesellschaft, SchmidGmbH, K€ongen, Germany; Fig. 1). For identification ofeukaryotic parasites the studies of Schenk (1858), Zopf

(1884), Sparrow (1960), Johnson and Sparrow (1961), Drebes(1966), Karling (1977) and Letcher and Powell (2012) wereused. For diatom species determination, samples of the slurry(3 9 5 mL) were prepared according to the method of Sch-rader (1974) and embedded in Naphrax (refractive index1.74, Northern Biological Supplies Ltd., Ipswich, UK). Forspecies identification the following literature was used: Cleve(1894, 1895), Hasle and Syvertsen (1996), Hendey (1964),Hustedt (1927–1966, 1939), Krammer and Lange-Bertalot(1986–91), K€utzing (1844, 1849), Schmidt et al. (1874–1959),Witkowski (1994) and Witkowski et al. (2000). All permanentsamples were examined una light microscope (Axiophot, CarlZeiss AG, Oberkochen, Germany) at 1,0009 magnification.Numerical processing of the identified diatom species wascarried out after classification (classes and orders only)according to the taxonomic nomenclature described inMedlin and Kaczmarska (2004).

Calculation of the infection rates. The percentage of infectedcells was calculated by dividing the number of infected cellsby the total number of host cells. The mean number of chytr-ids and oomycetes per cell (host) in the diatom populationwas also calculated, by dividing the total number of parasitesattached to algal cells by the total number of host cells, tonormalize the cell density among treatments. This value isreferred to as the mean intensity of infection (Holfeld 2000),reflecting the number of pathogens that succeed in attachingto their host.

Statistical analysis. Percentage infection and infectionintensity of eukaryotic parasites obtained from each treat-ment (untreated sediment samples as well as ultrasound-trea-ted and gradient-centrifuged ones) were compared usingone-way analysis of variance (ANOVA) followed by Tukey’stests. The data were checked for normality with Bartlett’s test(Bartlett 1947) and, where necessary, log transformed prior

FIG. 1. Illustration of the methods used for the collection of surface sediment samples in the Solth€orn tidal flat (A) as well as the sub-sampling scheme for each step of the preparation (B–D). * vials were prepared according to Schrader (1974).

EUKARYOTIC PATHOGENS INFECTING MPB DIATOMS 1011

TABLE1.

Overviewofthestainingmethodsusedduringthesurveysan

dtheireffectiven

essin

thedetectionofoomycetes

andch

ytridsin

thesamples.

Stainingagen

tAbbre-

viation

Supplier

Referen

ceincluding

stainingprotoco

lStainingspecificity

Visibilityofpathoge

nstructuresin

thisstudy

Acidfuch

sin

AF

F81

29;

Sigm

a-Aldrich

Dicksonet

al.20

03Stainske

ratin,cytoplasm

,an

dnuclei

Nuclei

from

active

cells,butnoem

pty

sporangia.

Tootimeco

nsuming,

dueto

lack

ofspecificity

Calco

FluorWhite

CW

1890

9;Sigm

a-Aldrich

Herth

1980

Bindsto

cellulose

andch

itin

incellwalls

Sporangiaan

dem

pty

sporangiavisible.Also

structuresofoomycetes

wereclearvisible.High

backstainingin

sedim

entsamples

Congo

red

CR

C67

67;

Sigm

a-Aldrich

Brigg

san

dBurgin

2004

Bindsto

chitin

incellwalls

Sporangiaan

dem

pty

sporangiavisible.

Nooomycetes

detected.Highbackstainingin

sedim

entsamples

FIT

C-conjugatedwheat

germ

agglutinin

WGA

FL-102

1;Biozol

Harris20

05Chitin-specificbindingprotein

Sporangiaan

dem

pty

sporangiavisible.

Nooomycetes


sedim

entsamples

Lactophen

ol-cotton

blue

LCB

PL.705

4;Pro-Lab

Diagn

ostics

Leck19

99Cottonbluestainsthech

itin

inthefungalcellwalls

Sporangiaan

dem

pty

sporangiavisible.

Nooomycetes


sedim

entsamples

Lactophen

ol-P

icric

AcidSo

lution

LPA

4150

6;Sigm

a-Aldrich

Flemingan

dSm

ith19

44Picricacid

bindsto

chitin-chitosanco

mplexe

sNoch

ytridsan

doomycetes

weredetected.

Highbackstainingin

sedim

entsamples

Mayer’s

Mucicarm

ine

StainSo

lution

MMS

4132

5;Sigm

a-Aldrich

Walsh

andJass

2000

Stainforcarbohydrates

Notspecific,

stained

allorgan

ismsco

ntaining

carbohydrates.Highbackstainingin

sedim

ent

samples

Neu

tral

red

NR

N28

89;

Sigm

a-Aldrich

Vierheiliget

al.20

05Stainslysosomes

Noem

pty

sporangiavisible.Tootimeco

nsuming,

dueto

lack

ofspecificity

Propidium

iodide

PI

P48

64;

Sigm

a-Aldrich

Riccardian

dNicoletti20

06DNAstain;usedas

anuclear

counterstain

Noem

pty

sporangiavisible.Tootimeco

nsuming,

dueto

lack

ofspecificity

Trypan

Blue

TB

T81

54;

Sigm

a-Aldrich

Phillipsan

dHayman

1970

Bindsto

chitin

incellwalls

Sporangiaan

dem

pty

sporangiavisible.

Nooomycetes


sedim

entsamples


to analysis. A P-value of <0.05 was considered as significant.All tests were performed with the program SPSS StatisticalSoftwareTM, version 11.5 (Edinburgh, UK).

RESULTS AND DISCUSSION

Sediment sample pre-treatment and stainingmethods. Initially, all ten staining methods used incombination with the untreated sediment sampleswere more or less ineffective, being not specificenough and/or giving high backscattering effectswith the sediment matrix (recovery rates only up to3.9%, Fig. 2; Table 1). Samples stained with Mayer’sMucicarmine stain (MMS) showed positive colorreactions with the exopolysaccharides (polymericcarbohydrates) originating from the benthic dia-toms and the sediment matrix, whereas propidiumiodide (PI), neutral red (NR) or acid fuchsin (AF)stained only all living organisms which were presentin the unpreserved sediment samples (e.g., algae,bacteria, cilliates, heterotrophic nanoflagellates).Conversely, empty sporangia and inactive cells, suchas those after an infection, were not stained by thesemethods. In particular, staining agents specific forchitin and its polymers such as CW, Congo red(CR), Lactophenol-cotton blue (LCB), Trypan Blue(TB) and WGA produced a manifold of false posi-tive signals with the fine and muddy sediments

(probably due to the presence of chitin in benthicdetritus). According to the manufacturer’s descrip-tion CW is a non-specific fluorochrome that bindsto cellulose and chitin in cell walls, whereas WGA isa sugar-binding protein (lectin) that binds only to aseries of consecutive N-acetylglucosamine residues(Roth 1978). In general, chitin rivals cellulose asthe most abundant biopolymer in nature (Menget al. 2012) and planktonic crustaceans havebeen considered the most significant source of chi-tin in the marine environment (Souza et al. 2011).In addition, copepod fecal pellets are encased inchitin (Yoshikoshi and Ko 1988) and several com-mon phytoplankton genera of diatoms, such asThalassiosira Cleve and Skeletonema Greville, producechitin as a significant portion of their biomass (upto 33%, Smucker and Dawson 1986). According toBoyer (1994) much of the chitin found in oceans israpidly degraded while in suspension, but some isalso incorporated into sediments (up to 20%,depending on the sediment composition). Thus it ishighly likely that the high backscattering effectsobserved in this study during the use of untreatedsurface sediment samples in combination with chi-tin-specific staining agents were caused by accumu-lated chitin residues originating from theplanktonic realm.The further pre-treatment of the sediment sam-

ples using ultrasound and gradient centrifugationresulted in an increase in the visualization of para-sitic structures (rhizoidal systems, thalli and sporan-gia), due to elimination of the backscatteringeffects. Both methods, ultrasound for the mechani-cal disruption of diatom-sand agglomerates as wellas density gradient centrifugation for the separationof sediment grains from the benthic diatom bio-mass, have already been applied by several scientiststo gain information about the community composi-tion of epipelic and epipsammic diatom taxa inhab-iting intertidal sediment surfaces (e.g., Admiraal1984, M�el�eder et al. 2005, Scholz 2014). In contrastto these protocols, only short ultrasound pulses(3 9 2 s) were used in this study, to conserve thefine structures of the eukaryotic parasites (thalli andattached sporangia), which were found to bedeformed or destroyed by pulses longer than 10 s(data not shown).Generally, the highest numbers of epi- and endo-

biotic parasites infecting benthic diatoms wererecorded by the use of CW (up to 46% and 68%higher in comparison to the untreated sedimentsamples, ANOVA: F1,90 = 32.9, P < 0.0001, Fig. 2),whereas the stains FITC-WGA, CR, LCB and TBshowed significantly lower recovery rates (up to 43%lower than the CW stained samples, ANOVA:F1,15 = 39.8, P < 0.0001). In addition, the high num-bers of presumed infections using the CW stain canbe explained by the counting of both groups: chytr-ids and oomycetes, whereas FITC-WGA, CR, LCBand TB seemed to be only sensitive for the chitin

FIG. 2. Comparison of autofluorescence counting and the tenstaining methods used during three sampling events foruntreated as well as pre-treated surface sediment samples fromthe Solth€orn tidal flat, presenting the prevalence (= ratio ofinfected cells). The individual staining methods are given inTable 1. Considered were the main epi- and endobiotic eukary-otic parasite structures such as thalli, sporangia, and rhizoidal sys-tems. Mean values (� SD), based on triplicate assays and thesamplings of one station from June to July 2012, are given.


containing cell walls of chytrids. Conversely, the dif-ferences in the susceptibility of the staining agentsused in this study in conjunction with the composi-tion of chytrid and oomycete cell walls (chitin vs.cellulose) could be the crucial factor for the nondetection of oomycetes in the untreated as well as

pre-treated sediment samples, suggesting the lack ofchitin in the oomycetes found during the presentshort-term monitoring.Many actinomycetes and fungi are known to fluo-

resce in UV (e.g., Rost 1995). Particularly underviolet excitation, weak to moderate fluorescence in

TABLE 2. Total diatom abundances (classes and orders according to the taxonomic nomenclature described in Medlin andKaczmarska 2004) and number of epibiotic and endobiotic eukaryotic pathogens. Data were obtained from overview coun-tings in Uterm€ohl chambers, using CalcoFluor White as staining agent. Mean values (� SD) for all subsamples are given(n = 9).

Diatoms/Sampling date June 16, 2012 July 01, 2012 July 14, 2012 Magnitude

Class: Coscinodiscophyceae 0.14 � 0.05 0.12 � 0.1 0.19 � 0.05 [103 cells � cm�2]Class: Mediophyceae 0.8 � 0.1 0.3 � 0.1 1.0 � 0.4 [103 cells � cm�2]Class: Bacillariophyceae

Order: Achnanthales 2.3 � 1.1 2.4 � 1.2 2.1 � 1.5 [103 cells � cm�2]Order: Bacillariales 1.3 � 1.0 0.8 � 0.4 1.0 � 0.6 [103 cells � cm�2]Order: Naviculales 3.6 � 1.1 3.1 � 0.8 2.8 � 0.9 [104 cells � cm�2]Order: Rhaphoneidales 0.3 � 0.01 0.23 � 0.01 0.42 � 0.02 [102 cells � cm�2]Order: Striatellales 0.42 � 0.02 0.38 � 0.01 0.59 � 0.02 [102 cells � cm�2]Order: Thalassiophysales 1.4 � 0.4 1.2 � 0.5 1.1 � 0.6 [103 cells � cm�2]

∑ 4.21 � 3.8 3.71 � 3.1 3.35 � 4.1 [104 cells � cm�2]Pathogens

Epibiotic 9.2 � 0.2 12.1 � 0.1 19.4 � 0.6 [102 cells � cm�2]Endobiotic 3.8 � 0.2 4.2 � 0.3 2.2 � 0.3 [cells � cm�2]

∑ 9.23 � 0.4 12.15 � 0.4 19.47 � 0.9 [102 cells � cm�2]Infection rates 2.2% 3.4% 5.8%

TABLE 3. Morphological characteristics of the epi- and endobiotic eukaryotic pathogens obtained from analysis of perma-nent slides, using sediment samples collected in the Solth€orn tidal flat in June and July 2012.

Pathogen Morphological characteristics Suggestions

Chytridiomycota (monocentric, holocarpic or eucarpic, epibiotic)Type I Sporangium: sessile, ovoid or spherical (globose), 14–25 lm

high, 14–26 lm diameter; wall thin, and smooth, colorless,double-contoured, 1–1.5 lm thick, with a broad apical or subapical papilla. Rhizoid: simple, short, thick, and peg-like,projecting only slightly beyond the inner part of the host wall

Rhizophydium Schenk (1858), Sparrow (1960),section 1 in Letcher and Powell (2012);similarities with R. brevipes var. marinum(but sporangia seems to be too small); restingspores not observed

Type II Sporangium: sessile, ovoid-obpyriform or urceolate (emptysporangia), 5–11 lm high by 5–9 lm in diameter; thick walled(2.5–3 lm), wall smooth, colorless; Rhizoid: rhizoidsnot observed

Rhizophydium Schenk (1858), Sparrow (1960),section 2 in Letcher and Powell (2012); restingspores not observed

Type III Sporangium on a short thick walled, rigid stalk(extrametrical, 1.3–2.5 lm long), ovate or ellipsoid, 12–15 lmlong by 9–11 lm high by 8–9 lm wide; without rhizoids,usually with a spherical “oil” drop like spot (3–4 lm),gregarious

Chytridium sp. Johnson and Sparrow (1961),according to Sparrow (1960) similarities withC. chlorobotrytis Fott with exception of stalk;resting spores not observed

Type IV Sporangium on a thin walled, flexible stalk (extrametrical,5–12.5 lm long), ovoid, obpyriform or spherical, 9.5–15.5 lmlong by 9–15.5 lm in diameter, with a thin, smooth, colorlesswall; gregarious, rhizoidal system short, delicate, and sparselybranched

Chytridium sp. Sparrow (1960), Johnson andSparrow (1961); resting spores not observed

Type V Sporangium on a flexible stalk (extrametrical, 12–44 lmlong), globose, 20–35 lm in diameter, hyaline, smooth,somewhat thick-walled (2–3 lm thick) solitary; rhizoid simple,short, thick, and peglike, projecting only slightly beyond theinner face of the host wall

Chytridium or Rhizophydium (Sparrow 1960),in general both groups include species withstalks (Sparrow 1960, Johnson and Sparrow1961, Letcher and Powell 2012); restingspores not observed

Oomycota (monocentric, holocarpic, endobiotic)Type I Sporangium single-celled, non-branched, inoperculate,

spherical, lenticular 26–40 lm long by 20–35 lm in diameter,with a thin colorless wall, discharge tubes from one to five,broadly conical, 8–10 lm long by 9–12 lm in diameter

Ectrogella Zopf (1884) or Olpiopsis sp. (!), becauseboth are known to form single-celled,non-branched endiobiotic sporangia in the hostcytoplasm (Sparrow 1960), the two genera differonly in spore morphology

Type II Thallus tubular, very thin-walled, 170–350 lm long by5–7.5 lm in diameter, infection tube occasionally persistent

Lagenisma Drebes (1966), according to Sparrow(1960) similarities to Lagenidium sp.


fruiting bodies has been observed in several fungalspecies, suggesting that autofluorescence is a com-mon feature of these taxa (Rost 1995). While forsome species fluorescent substances such as orella-nine (e.g., Cortinarius orellanus, Keller-Dilitz et al.1985) or leprocybin (subgenus Leprocybe, Kopanskiet al. 1982) are described, no references about suchsubstances were found for chytrids and oomycetes.However, due to the fact that autofluorescence wasused in previous investigations regarding the MPBcomposition in the Solth€orn tidal flat (Scholz and

Liebezeit 2012) it was also tested in connection tothe screening for eukaryotic parasites in this study.Here, autofluorescence yielded only poor results inall cases, which might be caused by the general lackof fluorescing structures and/or the damage anddenaturation of chloroplasts and fluorescing sub-stances, respectively, during the pre-treatments.A clear disadvantage of the pre-treatments was the

loss of free swimming zoospores during the prepara-tion steps, which are in most cases necessary for thespecies identification of chytrids and oomycetes

FIG. 3. Examples of diatomsinfected by different chytrids foundin the Solth€orn tidal flat. Sampleswere prepared as described in themethods & material part, using themodified CW method. Black barsindicate 10 lm. Pictures A, B, E andF were taken under fluorescencelight, whereas for C and D usualvisible light was used. (A)Cylindrotheca closterium/chytrid typeII; (B) Amphora exigua/chytrid typeIV; (C) Navicula digitoradiata/chytridtype III; (D) Achnanthes brevipes/chytrid type V; (E) Navicula gregaria/chytrid type I; (F) Diploneis didyma/chytrid type II.


(Sparrow 1960). An important aspect of the presentapproach is the use of settling chambers for detect-ing eukaryotic pathogens according to the classicalmethod of Uterm€ohl (1958). Here, the identificationof species was in most cases impossible. Similarresults were obtained by Rasconi et al. (2009), whofound that the main reason for this is generally thatstaining directly in the Uterm€ohl chamber resultedin very poor quality of parasite visualization. How-ever, the use of settling chambers in this study wasreduced to overview counts, distinguishing onlybetween epi- and endobiotic parasites (Table 2),while for identification purposes permanent slideswere prepared (Table 3; Fig. 3). In this context, theuse of polycarbonate filters (0.6 lm pore-size), whichare shown by Rasconi et al. (2009) to be practical forfreshwater phytoplankton samples, may be also use-ful in future investigations of MPB samples after apre-treatment such as applied in this study.Total benthic diatom community and infection rates.

The total diatom abundance did not vary signifi-cantly during the short-term monitoring from Juneto July 2012 (P > 0.05), decreasing only slightly overthe three sampling events (difference from 16 Juneto 14 July, 2012 = 7.6%, ANOVA: F1,16 = 34.7,P < 0.0001, Table 2). The data obtained from theoverview countings, using Uterm€ohl chambers incombination with the CW stain, revealed the Navi-culales as the most abundant group within the ben-thic diatom community of the Solth€orn tidal flat(up to 85.3% of the total diatom abundances). Gen-erally, the dominance of small species belonging tothe genus Navicula spp. appears to be an importantfeature of European intertidal MPB (e.g., Admiraalet al. 1984, Underwood 1994, Hamels et al. 1998).

Besides the dominance of the representatives ofthe Naviculales, the Achnanthales were identified as

the second most abundant order in the benthic dia-tom community (6.1%), whereas the Thalassiophy-sales (3.3%) and Bacillariales (2.8%) representedthe third and fourth most abundant group of taxa,respectively. All other groups had significantly lowertotal abundances and can be regarded as minorgroups of the MPB diatom communities of the Sol-th€orn tidal flat during this short sampling period.Only few diatoms were found to be infected by

eukaryotic pathogens, with the highest prevalencerecorded in mid July 2012 with 5.8% of the overallbenthic diatom community (Table 2). At all threedates of the survey, the numbers of epibiotic para-sites exceeded the occurrence of endobiotic ones byfar, ranging from 99.6% to 99.8% in June and July2012, respectively. Generally, chytrids are frequentlyfound to cause significant algal mortality in freshwa-ter plankton, where during epidemics more than90% of the host population may be infected (e.g.,Van Donk and Ringelberg 1983, Kagami et al.2006). Regarding the impact of oomycetes on theirhost population, it was reported for instance that anapprox. 13% infection prevalence in a natural popu-lation of Coscinodiscus was caused by L. coscinodisciDrebes in the Weser estuary of northern Germany(Raghukumar 1996).Epi- and endobiotic infections of benthic diatoms. Due

to the methods used, identification of epi- and endo-biotic eukaryotic pathogens was only possible atgenus level. Generally, the species identification ofeukaryotic parasites in environmental samples is verydifficult as they rarely show species-specificmorphological structures (Miller 1968) and observa-tions in samples obtained from a habitat generally donot allow observation of all phases of their full lifecycle. Most of these pathogens have not been estab-lished in pure culture yet where they could be studiedextensively under controlled and variable conditionsto reveal life cycle transitions. In addition, differencesbetween the morphology of in situ observations andspecies in culture have been found for some repre-sentatives of the Chytridiomycota (e.g., Chen andChien 1996), which can be explained, for example,by nutrient effects (e.g., Hasija and Miller 1971, Chenand Chien 1996). These environmental influences onmorphology further complicate the identification ofparasitic species in environmental samples.Overall five representatives of the Chytridiomycota

were distinguished according to their rhizoidal sys-tem and epibiotic sporangia (Table 2; Fig. 3). Thediatom taxa infected by chytrids comprised repre-sentatives of the Bacillariales, Naviculales, Achnant-hales and Thalassiophysales (Fig. 3). Specifically,Navicula digitoradiata (Gregory) Ralfs (Fig. 3C) wasinfected by chytrid type III, whereas two otherspecies of this order were found to be infected bychytrid type I (N. gregaria Donkin, Fig. 3E) and II(Diploneis didyma Ehrenberg, Fig. 3F). Additionally,chytrid type II was also observed on cells of Cylindrot-heca closterium (Ehrenberg) Reiman & Lewin

FIG. 4. Examples of diatoms infected by different oomycetesfound in the Solth€orn tidal flat. (A) Lagenisma Drebes (host: Cos-cinodiscus radiatus Ehrenberg) and (B) Ectrogella Zopf (host: Gyr-osigma peisonis [Grunow] Hustedt). Black bars indicate in (A) 30lm and (B) 15 lm.


(Fig. 3A) a representative of the Bacillariales. Fur-thermore, one representative of the Achnanthales(Achnanthes brevipes Agardh, Fig. 3D) was infected bychytrid type V, while one representative of the Tha-lassiophysales (Amphora exigua Gregory, Fig. 3B) wasinfected by type IV. It has to be pointed out, how-ever, that the taxonomic identity of these taxa haveto be revised (especially when molecular methodswill become available) considering recent studies onwhat are likely closely related pathogens in brownalgae (K€upper et al. 2006).

In addition, microscopic analysis of the endobioticthalli and sporangia, using permanent slides,revealed two representatives of the Oomycota (cf.Table 2, Fig. 4), Lagenisma Drebes (Lagenismatales,host: Coscinodiscus radiatus Ehrenberg, Coscinodisco-phyceae, Fig. 4A) and Ectrogella Zopf (Saprolegniales,hosts: Dimeregramma minor (Gregory) Ralfs in Prit-chard, Mediophyceae and Gyrosigma peisonis (Gru-now) Hustedt, Naviculales, Fig. 4B), respectively.

CONCLUSIONS

The combination of staining with CW and pre-treatment of sediment samples using short ultra-sound pulses and density gradient centrifugation,gave the best results in screening for eukaryoticpathogens in the benthic marine realm. Althoughthe Uterm€ohl method prevents taxonomic identifica-tion of the parasites, it is at least a useful tool to quan-tify infection rates of eukaryotic pathogens in thetotal microphytobenthic community by differentia-tion between epi- and endobiotic ones. Microscopicanalysis of permanent slides revealed the presence ofat least two oomycetes and five different chytrids.

We are grateful to Elisabeth Jaskulska and Dipl. Ing. ChristineWagener for their kind cooperation during the samplingcampaign. The MASTS pooling initiative (Marine Alliance forScience and Technology for Scotland, funded by the ScottishFunding Council and contributing institutions; grant refer-ence HR09011) is gratefully acknowledged.

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