Parasitic infections - CSIRO Publishing

Volume 37 Number 1 March 2016Volume 37 Number 1 March 2016

Parasitic infections

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Volume 37 Number 1 March 2016

ContentsFrom the Editorial Team 2

Jo, Ian, and Hayley Macreadie

Guest Editorial 3Parasitic infections: overlooked, under-diagnosed and under-researched 3

Harsha Sheorey and Richard S Bradbury

In Focus 4The laboratory diagnosis of Strongyloides stercoralis 4

Matthew R Watts, Gemma Robertson and Richard S Bradbury

Under the Microscope 10Current WHO protocols for mass drug administration in helminth control 10

Richard S Bradbury and Patricia M Graves

Zoonotic tissue parasites of Australian wildlife 12

David M Spratt

Assessing enteric helminths in refugees, asylum seekers and new migrants 15

Sarah Hanieh, Norbert Ryan and Beverley-Ann Biggs

Free living amoebae and human disease 20

Evan Bursle and Jennifer Robson

Tapeworm cysts in the brain: can we prevent it happening? 25

Marshall Lightowlers

Seafood-borne parasitic diseases in Australia: how much

do we know about them? 27

Shokoofeh Shamsi

Children, snails and worms: the Brachylaima cribbi story 30

Andrew R Butcher

Malaria: global challenges for malaria eradication 34

Graham Brown and Stephen Rogerson

Plasmodium knowlesi: an update 39

Balbir Singh

Diagnosis of human taeniasis 43

Abdul Jabbar, Charles Gauci and Marshall W Lightowlers

Lab Report 46Protozoa PCR: boon or bane 46

Colin Pham and Harsha Sheorey

ASM Affairs 50FT020 Water Microbiology Australian Standards Committee 50

2016 ASM Communication Ambassador Program 50

Stopping dengue: recent advances and new challenges 51

MICROBIOLOGY AUSTRALIA • MARCH 2016 1

Dr Gary Lum Dr John MerlinoProf. Wieland MeyerProf. William RawlinsonDr Paul SelleckDr David SmithMs Helen SmithDr Jack Wang

The Australian Societyfor Microbiology Inc.9/397 Smith StreetFitzroy, Vic. 3065Tel: 1300 656 423Fax: 03 9329 1777Email: [email protected] 24 065 463 274

For Microbiology Australiacorrespondence, see address below.

Editorial teamProf. Ian Macreadie, Mrs Jo Macreadieand Mrs Hayley Macreadie

Editorial BoardDr Chris Burke (Chair)Prof. Mary BartonProf. Linda BlackallProf. Sharon ChenProf. Peter ColoeDr Narelle FeganDr Geoff HoggProf. Jonathan IredellDr I

.pek Kurtböke

Subscription ratesCurrent subscription rates are availablefrom the ASM Melbourne offi ce.

Editorial correspondenceProf. Ian Macreadie/Mrs Jo MacreadieTel: 0402 564 308 (Ian)Email: [email protected]

Published four times a year in print and open access online by

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ISSN 1324-4272eISSN 2201-9189

While reasonable effort has been made to ensure the accuracy of the content, the Australian Society for Microbiology, CSIRO, and CSIRO Publishing accept no responsibility for any loss or damage from the direct or indirect use of or reliance on the content. The opinions expressed in articles, letters, and advertisements in Microbiology Australia are not necessarily those of the Australian Society for Microbiology, the Editorial Board, CSIRO, and CSIRO Publishing. Cover image: Protoscolex of Echinococcus granulosus, showing row of hooklets; obtained from liver

aspirate of a case of hydatid cyst. Courtesy of Associate Professor Rob Baird, Royal Darwin Hospital.

Welcome to the first issue of Microbiology Australia for 2016.

As members of The Australian Society for Microbiology know, our

electronic issues are freely available. To be notified of a new issue go

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register to get alerts when each issue becomes available.

ASM members receive print copies by mail if they have opted to

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print copies. Additional copies are often made for special issues.

For example, ~1000 copies were made for delegates of the ISHAM

conference in Melbourne last year. The print copies are highly

valued by many members and are great to share with others.

The online issues are highly accessed as a reference and learning

resource. Individual articles as well as entire issues can be readily

downloaded from the above website.

Themes of issues and Guest Editors are chosen by the Editorial

Board whomeet by teleconference five times each year. The Board

always values input from ASMmembers for future issues. All articles

are peer-reviewed and invited by theGuest Editorswho have a deep

understanding of the themes they present. Themes are chosen to

be topical and of interest to ASM members: authors write to reach

a broad audience.

On occasions urgent updates are needed and such updates are

presented asHot Topics. However, we are aware that there is a need

to have an occasional non-themed issue. Please contact us if you

have suggestions for articles that will have broad interest to ASM

members.

We are grateful for the support the Editorial Board and for Guest

Editors who are experts in their fields. Guest Editors present the

latest knowledge in their field through the selection of contributors

who write compact articles that are readable and informative to the

ASM’s diverse membership. We also thank the reviewers, who

provide valuable comments on each contribution. They also have

expertise in the articles they review, and they provide valuable

comments to contributors to ensure the articles are appropriately

presented. We also thank the authors for their contributions. They

provide a great amount of current information about their field and

seek to educate us all in the exciting developments ofmicrobiology.

We thank The Australian Society for Microbiology and those who

lead it. In thechallengingworldofpublishing ithas takenboldness to

enableMicrobiology Australia to be produced as a publication that

is freely available to all. We know from the download data provided

byCSIROPublishing, thatMicrobiology Australia is highly accessed

around the world, and that the articles continue as an ongoing

resource many years after their first publication. Articles published

more than a decade ago still enjoy thousands of downloads every

year. There is no doubt that they serve as resource material for

teachers, students and those seeking reliable scientific information

from experts.

We trust that you enjoy your reading of this year’s issues, which will

cover:

* Parasitic Infections* Education to enhance microbiology graduate employability* Diseases of Aquaculture* Microbiology of Travel [a special joint issue with The MicrobiologySociety in the UK].

Jo, Ian, and Hayley Macreadie

Access to Microbiology Australia

Online early articles: Articles appear on the Microbiology Australia website (http://microbiology.publish.csiro.au/) when authors have approved the pdf of their article.

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Print issue: ASM members are welcome to receive the print version of Microbiology Australiawithout charge. To receive the print version you need to notify the ASM National Office (http://www.theasm.org.au/).

2 10.1071/MA16001 MICROBIOLOGY AUSTRALIA * MARCH 2016

From theEditorial Team

Parasitic infections: overlooked, under-diagnosedand under-researched

Harsha Sheorey

Microbiology DepartmentSt Vincent’s Hospital MelbourneFitzroy, Vic., AustraliaEmail: [email protected]

Richard S Bradbury

School of Medical and AppliedSciencesCentral Queensland UniversityRockhampton, Qld, AustraliaEmail: [email protected]

ProfessorGeorgeNelson (1924–2009) once stated that, ‘Parasitology

is the preserve of the diagnostically destitute’. Little has changed to

thisday,withpotentially relevantparasiticcausesof illnessesoftennot

being considered early in the differential diagnoses of clinical pre-

sentations. Parasitic infections are sometimes overlooked as causes

of morbidity and (in some cases) mortality in both the medical and

veterinary fields. In Australia there remain significant problems

associated with giardiasis, cryptosporidiosis, strongyloidiasis and

other parasitic diseases, particularly in remote, underserved and

tropical regionsof the country and also in the immuno-compromised

individuals (HIV, immunosuppressive drugs etc.). The burden of

many parasitic diseases is greater in tropical and sub-tropical areas

of non-industrialised countries. With increasingly adventurous travel

and dining, increasing numbers of Australians returning from travel

overseas with added souvenirs of common or exotic parasitoses

every year and refugees and migrants arriving in Australia, these

infections are becoming increasingly important.

Recent advances in diagnostic techniques for the detection of

parasitic infections have revolutionised how we undertake such

diagnostic investigations. These methodologies often provide

greater sensitivity as well as in some cases providing additional

epidemiological data. As we increasingly move towards the use of

molecular methodologies and away from traditional morphological

diagnosis, newchallenges have emerged for clinicians, veterinarians

and laboratory staff. A focus of several articles in this edition is

consideration of the advantages and disadvantages of these new

methods, in what circumstances they are best applied and how the

results of such investigations should be best interpreted. Molecular

methods are not without potential sources of error, hence in some

situations, morphological and serological techniques still remain

relevant.

This edition also considers zoonotic parasitic infections, the treat-

ment of parasitic infections, both from the current WHO recom-

mendations formass drug administration inhighly endemic settings

to the rise of resistance to anti-parasitic agents in protozoa such as

malarial parasites and Giardia intestinalis. Whilst antimicrobial

resistance is greatly investigated in bacterial infections, its emer-

genceandprevalence inparasitic infectionsofhumanandveterinary

importance will require further investigation and attention in the

future.

We hope that this edition of Microbiology Australia will update

knowledge andserve to informall our readersof the importance and

relevance of parasitic disease. Whether one is involved in medical,

veterinary, food, environmental or other microbiological work, it is

likely that aspects of your work will at some stage involve this

important and sometimesneglectedfieldof our scientificdiscipline.

As guest editors, we are grateful and excited to be involved in the

planning and execution of this edition. We would like to thank the

editorial staff and all of the authors and reviewers who have kindly

contributed their time and expertise into the preparation of this

edition. We hope that all members of the society will find it helpful,

interesting and that it may spark the interest of many into this most

fascinating and under-researched area.

Biographies

The biography for Dr Harsha Sheorey is on page 49.

The biography for Dr Richard Bradbury is on page 9.

GuestEditorial

MICROBIOLOGY AUSTRALIA * MARCH 2016 10.1071/MA16002 3

The laboratory diagnosis of Strongyloidesstercoralis

Matthew R WattsA,*, Gemma RobertsonB,* and Richard S BradburyC,D

ACentre for Infectious Diseases and Microbiology, Pathology West – ICMPR, and Marie Bashir Institute, University of Sydney, Westmead Hospital,Westmead, Sydney, NSW, Australia, Tel: +61 2 9845 6255, Email: [email protected]

BMelbourne Pathology, Collingwood and James Cook University, Tel: +61 3 9287 7700, Email: [email protected]

CSchool of Medical and Applied Sciences, Central Queensland University, Rockhampton, Qld, Australia

DCorresponding author. Email: [email protected]

It is estimated that over 30million people worldwide are

infected by the nematode, Strongyloides stercoralis1. It is

endemic in sub-tropical and tropical parts of Australia, with

high rates of infection documented in some indigenous

communities2. Due to the potential for chronic autoinfec-

tion, that may persist for decades, migration leads to the

presence of the infection in non-endemic areas1. Transmis-

sion tohumans isgenerally throughthepenetrationof larvae

through the skin, following contact with faecally contami-

nated soil1. Disease severity ranges from asymptomatic

chronic carriage to an overwhelming illness, where large

numbers spread throughout the body, usually triggered by

immunosuppression1.

Clinicians areadvised toconsider strongyloidiasis inpatientsprior to

immunosuppression, or with indicative symptoms, if there is a

history of probable exposure in an endemic area, regardless of the

elapsed time since exposure3,4. Eosinophilia is not an accurate

marker of strongyloidiasis, with a retrospective study finding that

only a quarter of patients with Strongyloides infection had a raised

eosinophil count5. The detection of strongyloidiasis is optimised by

appropriate test ordering, clinical notes, specimen transportation,

and processing by the receiving laboratory.

The gold standard for the diagnosis of strongyloidiasis is the

morphological identification of larvae in stool, tissue biopsies, and

other clinical specimens such as bronchoalveolar lavage. However,

in chronic infections, detection canbe limited by low larval output in

stool, leading to false negative results6. Consequently, in validation

studies for serological and nucleic acid tests there is a tendency

to define heavier infections as ‘true positives’. This affects serolog-

ical cut-offs, measurements of sensitivity and specificity, and

positive and negative predictive values7. Recognition of these

limitations is important for the interpretation of negative diagnostic

test results, where clinical suspicion remains. Here, we will give an

overview of currently available conventional and molecular tests

for the diagnosis of strongyloidiasis.

Stool microscopy and culture methods

Specimen transport and storage have amajor impact on the efficacy

of culture techniques in the laboratory diagnosis of S. stercoralis

from faecal samples. Fresh, unrefrigerated samples should be

delivered to the laboratory for culture as soon after collection as

possible, as the viability of larvae decreases incrementally with

storage at 48C over a 72-h period8. Rhabditiform and filariform

larvae will be found along with free-living adults of S. stercoralis in

*These two authors contributed equally.

In Focus


older cultures (Figure 1). Larval stages must be differentiated from

those of hookworms, which may also be recovered.

Microscopic methodologies such as examination of Kato-Katz pre-

parations, FLOTAC, and formalin/ethyl acetate concentrates have a

low yield compared to culture9,10. Amodified formalin/ethyl acetate

method proposed by Anamnart et al. improved rates of detection9.

Overall, however, microscopic techniques alone are insensitive and

not sufficient for the exclusion of strongyloidiasis. In one of these

studies, though 30 of 254 participants were diagnosed with stron-

gyloidiasis by either agar plate culture (APC; Figure 2) or Baermann

culture techniques, no infections were identified by microscopy

using the Kato-Katz technique11.

APC is possibly the easiest culture to perform in the context of high

volume diagnostic testing. Results are available within two days,

although extended incubation up to four days increases yield8,11,12.

Two studies comparing 48 h APC with Baermann culture found an

improved recovery of S. stercoralis larvae inAPC11,12. Recovery rates

improve markedly with multiple stool cultures6,11,13

Serological diagnosis

Several tests for the serological diagnosis of strongyloidiasis have

been described, using both crude and recombinant antigens. Two

commercial ELISA kits employing somatic antigens are available

from BORDIER (Strongyloides ratti antigen) and IVD Research

(S. stercoralis antigen), respectively14. Recently, two recombinant

antigens (32 kD recombinant antigen, called NIE and S. stercoralis

immmunoreactive antigen, SsIR) have been employed for serolog-

ical testing in both ELISA and luciferase immunoprecipitation sys-

temassay (LIPS) platforms15. The reported sensitivity and specificity

of various serological platforms ranges from 56-100% and 29-100%,

dependent upon themethod, antigens, cut-offs, study populations,

and referencemethods employed16. Strongyloides serology using a

crude larval extract antigen was shown in one study to be less

sensitive for the diagnosis of returned travellers (73%) compared

topatientswhohave lived for anextendedperiod in anendemicarea

(98%)17. No definitive study of serological methods has been con-

ducted to date, and much of the available data is subject to flaws in

methodology, particularly the use ofmicroscopy only as a reference

(a) (b)

(c)

Figure 1. Life stages of Strongyloides stercoralis in agar plate culture: (a) rhabditiform larva; (b) a free-living adult male with filariform larva adjacent;and (c) a gravid, free-living adult female with filariform larva adjacent.

In Focus

MICROBIOLOGY AUSTRALIA * MARCH 2016 5

standard for positive specimens and varying S. stercoralis exposure

rates amongst tested serum groups16. A 2010 study with a reference

standard of a combination of three culture methods and sedimen-

tation concentration found NIE LIPS had a sensitivity of 97.8% (cut-

off 37.89 LU) in a study population with high endemicity but from

regions without filarial infection15. Lower sensitivity resulted when

testing the same samples by NIE ELISA(84%), NIE-SsIR LIPS (91.2)

and a S. stercoralis crude antigen extract ELISA (97%)18. All assays

tested showed 100% specificity in this study15. A more recent study

comparedan in-house crude S. stercoralisfilariform larvae immuno-

fluorescent antibody test (IFAT)with the ELISAs from IVDResearch,

Bordier, anda recombinant antigenNIEELISA andLIPS14. This study

used reference samples identified as positive by culture as well as

microscopy, and also a composite reference standard of concordant

results in at least three of five serological tests14. The in-house IFAT

was found to be the most sensitive (93.9%) when used in a test

subject groupwithnoknownpreviousexposure to S. stercoralis and

using the composite reference standard, whilst NIE LIPS was found

to be the most specific test (100%)14. Furthermore, when tested

against subjects with potential previous exposure and using the

composite reference standard, NIE LIPS was almost 100% specific

and 84.6% sensitive (cut off value 1388 LU)14. In testing against the

same sample group, the Bordier and IVD ELISAs maintained a high

specificity (almost 100%), but a lower sensitivity (70% and 79%,

respectively) and the NIE ELISA showed the highest specificity

(99%), but a low sensitivity (45%) (cut-off 76.5 U/mL)14.

Seroreversion following treatment of many, but not all, patients

was noted in a study using a Strongyloides ratti antigen ELISA3.

However, this effect is not universal and varies between studies3,17.

Immunosuppression was demonstrated to cause a reduction in

serological sensitivity (62% vs previously determined 92%), when

testing haematological patients on antineoplastic therapies18. NIE

LIPS did not cross-react with antigens from other parasites in the

study by Bisoffi et al., whereas IFAT and the two commercial ELISAs

did yield false positives, particularly from Mansonella perstans

infection14. Suchcross-reactionmaybedecreasedbypre-incubation

of serum in an extract of Onchocerca gutterosa16.

Nucleic acid tests

Nucleic acid tests complement non-molecular methodologies for

the diagnosis of S. stercoralis, and allow the use of refrigerated,

frozen, or preserved specimens11,19,20. This simplifies specimen

transportation, particularly where collection occurs some distance

from the testing laboratory, and there is no risk of laboratory-

acquired strongyloidiasis21. DNA extraction and amplification can

be performed within 1 day, however, laboratories may batch speci-

mens according to demand.

DNA extraction

It is important that DNA extractionmethods for stool specimens are

effective at removing the numerous nucleic acid test inhibitors in

stool22. A comparisonof5methodsofDNAextractiondemonstrated

that two column-basedmethodswere themost effective for the PCR

detection ofDNA from Strongyloides ratti that had been spiked into

human stool. These were the MoBio PowerSoil kit (MoBio Labora-

tories, Carlsbad, CA, USA) and a method based on modifications of

Figure 2. Larval tracks of Strongyloides stercoralis on Koga agar plate culture.

In Focus

6 MICROBIOLOGY AUSTRALIA * MARCH 2016

the QiaAmp Tissue kit (Qiagen, Hilden, Germany) by Verweij et al.,

which has been successfully automated23. The comparison

found that bead beating prior to the use of the NucliSens EasyMag

(BioMerieux, Marcy l’Etoile, France) was less effective, which indi-

cates the method of sample pretreatment prior to automated

extractionwill impact upon test sensitivity. Other investigators have

used a variety of different extractionmethods for StrongyloidesPCR,

including in-house methods, the Qiagen stool kit (unmodified and

modified), and the Nucleospin Soil kit (Macherey-Nagel, Duren,

Germany)10,24–30.

One of the inherent limitations of the molecular diagnosis of

S. stercoralis is the sampling error that can occur when relatively

small amounts are extracted in the context of low larval output6.

For example, 2g of stool can be used for agar plate culture, whereas

250mg of specimen is recommended for the MoBio PowerSoil

kit31. Methods that concentrate larger amounts of stool prior to

DNA extraction have the potential to increase test sensitivity, if

they remove inhibitors and retain larvae29.

PCR

Current PCR methods most commonly target one of four regions:

the 18S rRNA small subunit (SSU); the internal transcribed spacer

region 1 (ITS-1); the 28S rRNA gene; or the cyclooxygenase gene

(cox1)19,23–30,32–37. Published sensitivities and specificities for

Strongyloides PCR vary according to the reference methods and

are listed in Table 1. Themajority of Strongyloides PCR publications

have used a real-time method with primers and probe published

by Verweij et al.19. This has also allowed for the development of

multiplexed PCR10,30,34,38. Some studies evaluating the diagnostic

accuracy of these PCR methods have used both morphological

diagnosis and detection of PCR products as their reference stan-

dards, and are not reviewed here. Theirmethodology precludes the

calculation of sensitivity and specificity based on gold-standard,

according to an FDA Guidance39.

In the absence of a consistent gold standard in chronic infection,

positive nucleic acid test results, where conventional tests are

negative, may be due to greater sensitivity or false positive results6.

No PCR studies have reported false positive results when analytical

specificity has been tested using DNA extracted from bacteria,

viruses, fungi, protozoa, and other helminths19,23,24,27,29,30. Studies

have also assessed the specificity of the PCR products by sequence

analysis, with all finding 100% sequence homology with the target

sequence of S. stercoralis.24,25,27,29. Sitta et al. found a number of

false positives, using published genus and species-specific primers,

based on non-target sized bands on gel electrophoresis19,25. The

genus-specific primers amplified sequences that generated non-

target bands on electrophoresis in specimens that contained

Blastocytis and other helminths on microscopy, and the species-

specific primers amplified sequences that generated non-target

bands on electrophoresis in specimens positive for hookworm on

microscopy25. Similar accounts of cross-reactivity have not yet been

reported, so further data will be useful to monitor the specificity of

PCR in different populations.

Table 1. Sensitivity and specificity of stool PCR for human strongyloidiasis.

Reference Target Sensitivity Specificity Reference method

19 18S 61.0% 92.4% Coproculture; Baermann

32 18S 58.6%/96.6%A ND McMaster

26 18S 61.0% 92.7% APC; Baermann

34 18S 100% 100% Direct microscopy

23 18S 33.0% 99.0% Harada-Mori

25 18S 84.8%/78.8%A ND APC

10 18S 11.6%83.3%

90.6%96.2%

BaermannFLOTAC

29 18S 93.8% 86.5% FEAC; APC; Harada-Mori

36 18S 90.0% 85.7% APC

27 18S/cox1B

18S100%84.7%

91.6%95.8%

FEC; APC

AThe first value relates to a species-specific primer, the second to a genus-specific primer.BNested PCR. APC, agar plate culture; FEAC, formalin-ethyl acetate concentration; FEC, formalin-ether concentration.

In Focus


LAMP

Loop-mediated isothermal amplification (LAMP) is an additional

nucleic acid detection method. LAMP uses a DNA polymerase with

strand-displacement activity, so it doesn’t require the temperature

cycling of PCR, and can be performed with a simple source of

constant temperature such as a heating block40. LAMP has been

successfully applied in resource limited-settings for the detection

of pathogens40.

The Strongyloides LAMP assay uses primers that are genus specific

and bind to the 28S rRNA gene20. The reaction runs at 608C for

1 hour. Pre-heating of the reagents andDNA template to 958C, prior

to the addition of enzyme, increases the limit of detection and

eliminates the need to pre-heat the template and keep it at 48C20.

A novel use of Syto-82 dye (Life Technologies, Carlsbad, CA, USA)

enables the detection of positive results in real-time or visually on

completionof the reaction20. Analytical sensitivity and specificity are

comparable to PCR, according to themethodof Verweij et al.19,20,23.

When 28 human stool specimens that were microscopy and PCR

positive for S. stercoralis were tested with the LAMP method, 27

were positive20. The negative specimen had a high cycle threshold

(38.44) on PCR20. Further validation of the LAMP assay with clinical

specimens is currently in progress.

Conclusion

The diagnosis of strongyloidiasis can be made through the mor-

phological identification of larvae, usually in the stool, serological

testing, and nucleic acid tests. While each methodology has advan-

tages, there are limitations that need to be taken into account when

assessing the significance of negative test results. Often the most

important aspect of patient management is to consider the possi-

bility of S. stercoralis infection.

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Biographies

Matthew Watts is an Infectious Diseases Physician and Clinical

Microbiologist based at the Centre for Infectious Diseases and

Microbiology, PathologyWest-ICPMRand theMarieBashir Institute,

University of Sydney, Westmead Hospital. His interests include

parasitic and zoonotic infections.

Gemma Robertson is a final year microbiology trainee at Mel-

bourne Pathology. She has an interest in tropical medicine and

parasitology, andwill beundertaking aPhD tocontinueher research

into soil-transmitted helminthiases in Aboriginal communities.

Dr Richard Bradbury is an Australian Parasitologist with an

interest in all fields of parasitology. He was recently appointed as

the Team Lead in the Parasite Diagnostics and Biology Laboratory

of the Centers for Disease Control and Prevention in Atlanta, USA.

He is writing this work in both his personal capacity and in his

capacity as an adjunct academic at Central Queensland University.

In Focus


Current WHO protocols for mass drugadministration in helminth control

Richard S Bradbury

School of Medical and AppliedSciencesCentral Queensland UniversityRockhampton, Qld, AustraliaEmail: [email protected]

Patricia M Graves

College of Public HealthMedical and Veterinary SciencesDivision of Tropical Health andMedicineJames Cook UniversityCairns Qld, AustraliaEmail: [email protected]

Soil transmitted helminths (STH), comprising Ascaris,

Trichuris, Strongyloides and the hookworms remain a sig-

nificant cause ofmorbidity amongst people inmany parts of

the world, including Australia. Other important helminth

infections include lymphatic filariasis (LF), schistosomiasis

and onchocerciasis. Preventive chemotherapy (mass drug

administration [MDA]) campaigns are frequently conducted

for thesehelminth infections inendemicareas,but the target

population groups, duration of campaigns, cointerventions

(e.g. vector control) criteria for inclusion, drugs used and

doses of drugs differ.

The benefits of deworming individuals, especially children, who are

infected with soil-transmitted helminths and schistosomiasis,

include reduction in anaemia and improved growth. Rarer, but

more severe presentations, such as intestinal obstruction with

A. lumbricoides and rectal prolapse due to T. trichiura infection,

will also be reduced1. Treatment for filariasis and onchcocerciasis

in childhood will prevent the development of later severe conse-

quences of these diseases including lymphoedema, hydrocoele,

elephantiasis and blindness.

In situations ofmoderate to high endemicity, it has been considered

more efficient and cost-effective to treat the entire eligible popu-

lation in particular age groups or communities for these diseases,

rather thanfirst testing individuals todeterminewho is infected.The

goals of such MDA are morbidity control in some cases and inter-

ruption of transmission through vectors in others. For these rea-

sons, MDA for STH is usually undertaken for school age children,

while for schistosomiasis, MDA is performed either in children or

in eligible people of all ages, depending on the endemicity level.

For the insect borne helminths, onchocerciasis and LF, the goal is

transmission interruption and the entire community is eligible for

MDA.

The frequency of MDA for STH in school-age children is dependent

on the prevalence of infections in a given population (Table 1).

Current WHO protocols for STH control recommend MDA with a

single oral dose of albendazole (400mg), mebendazole (500mg) or

levamisole (80mg)1. Mebendazole is more effective than albenda-

zole for T. trichiura, whilst albendazole is slightly more effective

against hookworm than mebendazole. The two drugs have equally

Table 1. Current recommended regularity of mass drug administration(MDA) for helminth (STH) infections in school-ageA children (adaptedfrom WHO 2011)1.

PrevalenceB Regularity of MDA

For control of soil-transmitted helminth (STH) infections

�50% Twice per year, or every 4 months if at thehigh end of prevalence

�20 and <50% Once yearly

<20% Treat on case-by-case basis

For control of schistosomiasis

�50% Once per year, or every 4 months if at thehigh end of prevalence

�10 and <50%B Once every 2 years

<10% Treat on case-by-case basis

AUsually defined as children between 5 and 14 years of age.BAs determined by parasitological methods; cut-off is�30% if based only onquestionnaires for visible haematuria.

Under theMicroscope


high efficacy when used against Ascaris lumbricoides1. Despite

these differences, in practice when administered biennially over a

number of years, either drug used on its own is effective for overall

STH control1.

Onchocerciasis control and/or elimination requires annual or bian-

nual treatment with ivermectin for many years (up to 20 in some

cases), while lymphatic filariasis uses annual administration with

albendazole and either ivermectin or diethylcarbamazine (DEC)

with at least 65% population coverage for at least five years. Thus

ivermectin and/or albendazole administration may occur in some

communities annually or biannually as part of onchocerciasis or LF

elimination programs. Where this occurs, STH control programs

should be harmonised to ensure that there is a 6 month delay

between albendazole administrations1. In communities with en-

demic schistosomiasis, the addition of praziquantel (40mg/kg) is

recommended (Table 1). Reductions in the frequency of MDA may

be considered after 5–6 years of consistent >75% population cov-

erage, after testing of the residual prevalence of helminths in that

population. Such a decision is based on several factors, specific

details of which may be found in the World Health Organization

guide for managers of control programmes1.

Reliance on albendazole and mebendazole in WHO recommenda-

tions for MDA will result in a lower impact on the clearance of

Strongyloides stercoralis, for which ivermectin and thiabendazole

are more effective drugs2. Thiabendazole was discontinued in

Australia in 2003 anddue to the lower rate of side effects, ivermectin

has been recommended by some as the treatment of choice2. Due

to the auto-infective cycle of this helminth, some authors have

recommended re-treatment at one and two months to ensure

elimination3. Only one randomised trial has been performed thus

far, in which treatment twice at 2 weeks apart was found to have no

greater benefit than a single dose4. Both the authors of this paper

and others2 recommend further studies into the optimal dose

schedules for ivermectin in the control of Strongyloidiasis within

a larger cohort of participants.

Recently, there has been discussion about the outcomes and

optimal age range for MDA programs to control STH and more

evidence on the impacts is required5–7; this controversy is outside

the scope of this review. A novel concept of elimination of STH by

MDA ‘one village at a time’ rather than by wide-scale MDA has been

proposed for remote areas with low populations, such as many

remote Australian Aboriginal communities. This approach is cur-

rently being trialled in a remote area of the Solomon Islands8. It

advocates allowing individual communities to act as autonomous

units and to employ control options specifically tailored to the

geographic, cultural, economic, aetiological and environmental

factors influencing STH transmission in their own community8

(Figure 1).

Deworming of school-age children has been amainstay of helminth

control for many years. Discussion continues on the optimal meth-

od ofMDA for this purpose. In some cases, such as where high rates

of strongyloidiasis or onchocerciasis are present, the addition of

other drugs may be warranted. Further consideration of new ther-

apies, combination therapies, the reconsideration of use of the

use of ‘old’ anti-helminthic therapies have all been postulated as

mechanisms by which to improve absolute cure rates in MDA

programsand to reduce thepossibledevelopmentof antihelminthic

resistance9.Ways to improveMDAparticipation, aswell as paediatric

formulations of praziquantel for schistosomiasis prevention in pre-

school children may also be added to this list. The current WHO

protocols for MDA provide an important baseline guide to those

undertaking MDA for helminth control in endemic areas.

References1. World Health Organization (2011) Helminth control in school-age children:

a guide for managers of control programmes. 2nd edn. Geneva: World Health

Organization.

2. Bisoffi, Z. et al. (2011) Randomized clinical trial on ivermectin versus thiabendazole

for the treatment of strongyloidiasis. PLoS Negl. Trop. Dis. 5, e1254. doi:10.1371/

journal.pntd.0001254

3. Adams, M. et al. (2003) Strongyloidiasis: an issue in Aboriginal communities. Rural

Remote Health 3, 152.

4. Suputtamongkol, Y. et al. (2011) Efficacy and safety of single and double doses of

ivermectin versus 7-day high dose albendazole for chronic strongyloidiasis. PLoS

Negl. Trop. Dis. 5, e1044. doi:10.1371/journal.pntd.0001044

5. Anderson, R.M. et al. (2015) Should the goal for the treatment of soil transmitted

Helminth (STH) infections be changed from morbidity control in children to

community-wide transmission elimination? PLoS Negl. Trop. Dis. 9, e0003897.


6. Taylor-Robinson, D.C. et al. (2015)Deworming drugs for soil-transmitted intestinal

worms in children: effects on nutritional indicators, haemoglobin and school

Figure 1. A Solomon Islander researcher undertaking a community wideSTH prevalence survey as part of an integrated STH control program(photograph by Richard Bradbury).

Under theMicroscope


performance. Cochrane Database Syst. Rev. 7, CD000371. doi:10.1002/14651858.

CD000371.pub6

7. Hicks, J.H. et al. (2015) The case for mass treatment of intestinal helminths in

endemic areas PLoS Negl. Trop. Dis. 9, e0004214. doi:10.1371/journal.pntd.000

4214

8. Harrington, H. et al. (2015) Prevalence of soil transmitted helminths in remote

villages in East Kwaio, Solomon Islands. Western Pac. Surveill. Response J. 6, 1–8.

doi:10.5365/wpsar.2015.6.1.016

9. Savioli, L. (2014) Preventive anthelmintic chemotherapy – expanding the arma-

mentarium. N. Engl. J. Med. 370, 665–666. doi:10.1056/NEJMe1312403

Biographies

Dr Richard Bradbury is an Australian Parasitologist with an

interest in all fields of parasitology. He was recently appointed as

the Team Lead in the Parasite Diagnostics and Biology Laboratory

of the Centers for Disease Control and Prevention in Atlanta, USA.

He is writing this work in both his personal capacity and in his

capacity as an adjunct academic at Central Queensland University.

Dr Patricia Graves is a vector-borne disease epidemiologist with

research, field, laboratory and consulting experience in the control

and elimination of malaria and filariasis. She is Director of the

JCU/WHO Collaborating Centre for Control of LF, STH and other

NTDs in the Division of Tropical Health and Medicine, James

Cook University.

Zoonotic tissue parasites of Australian wildlife

David M Spratt

Australian National WildlifeCollectionNational Research CollectionsAustralia, CSIROGPO Box 1700Canberra, ACT 2601, AustraliaTel: +61 2 6242 1648Email: [email protected]

Increasing use of bushlands for recreational, commercial

and scientific activities fosters movement across the urban-

bushland interface. This may facilitate the transmission of

parasitic diseases from wildlife to humans (zoonoses). The

fashionable trend to consumption of game meats such as

feral pig and crocodile, and raw fish such as sushi, sashimi

and pickled herring has exacerbated the zoonotic potential

of parasites of wildlife.

Transmission from wildlife to humans

Angiostrongyliasis

Angiostrongylus cantonensis is a nematode parasite of the pulmo-

nary arteries and right ventricle ofRattus rattus andR. norvegicus in

Australia1. It is the causative agent of eosinophilic meningoenceph-

alitis, a zoonotic infection of humans. The life cycle includes an

obligatory period of larval development in terrestrial or aquatic

snails and slugs, and also may involve a range of paratenic or

transport hosts (freshwater prawns, land crabs, planarians, frogs,

lizards), which feed on gastropods. Rats become infected by ingest-

ing intermediate or paratenic hosts. In the rat, the nematode

undergoes an obligatory migration through the spinal column and

brain en route to the final site in the pulmonary arteries of the lungs.

Humans become infected by accidentally or deliberately eating

infected gastropods or paratenic hosts, or unwashed salad greens

containing these.Theparasitehasbeen reported fromdomestic and

zoo animals, mammalian and avian wildlife and humans in Brisbane

and Sydney2–4. The clinical signs of headache, vomiting, paralysis

and sometimes death are induced as a consequence of the oblig-

atory period of development of the parasite in the central nervous

system. This occurs in young children who deliberately or acciden-

tally ingest snails or slugs containing infective larvae5–7, or foolish

young adults who do so for a bet8,9.

Muspiceoidosis

Haycocknema perplexum is aminutemuspiceoid nematode living

as adults inside individual skeletal muscle cells of humans in

Australia10. Eight cases have been documented, 4 in Tasmania and

4 in north Queensland11–13 Gasser (personal communication).

Eight to twelve eggshatch inside theuterusof the female, develop to

third-stage infective larvae andburst fromthehead regionkilling the

adult, an efficient mechanism for auto-re-infection. Escaped larvae

invade uninfected muscle cells. The occurrence ofH. perplexum in

intramyofibres results in eosinophilic polymyositis but no reaction

within the invaded cell itself11. Progressive myopathy occurs and

infection becomes life threatening. Early human diagnosis by mus-

cle biopsy is imperative in cases of progressivemyopathy associated

with blood eosinophilia and elevated creatine kinase levels. Steroid

Under theMicroscope


treatment of patients exacerbates their infections to a life-threat-

ening illness and may delay diagnosis by masking key diagnostic

features. Treatment with albendazole, 400mg twice daily for 8–

9 weeks, is recommended12. Haycocknema perplexum is consid-

ered a zoonosis although the source of infection of humans –water,

soil, plants or animals – remains unknown.

Halicephalobus gingivalis

Halicephalobus gingivalis formerly known asMicronemadeletrix,

is a free-livingnematodeof soil,manure anddecayinghumusknown

to cause opportunistic infections, primarily in horses but also in

humans14.Themajorityof cases inhorseshavebeen fatal andusually

not diagnosed before necropsy. All human cases have involved fatal

meningoencephalitis including the first human case in Australia, a

74-year-old woman from Eyre Penninsula, South Australia15. In

tissues, only ova, larvae and adult females are seen, reproduction

in the parasitic phase presumed to be by parthenogenesis16. It is not

known how H. gingivalis infects humans or horses although expo-

sure through an oromaxillary route may explain common neuro-

logical involvement15.

Transmission from domestic animals to humans

and wildlife

Toxoplasmosis

Felids, domestic cats in particular, are the only definitive host of the

obligate intracellular protozoan parasite, Toxoplasma gondii17.

Most species of mammals and birds are susceptible to infection

and may act as intermediate hosts. Infection is usually systemic

resulting in a short period of rapid multiplication in various tissues

followed by the establishment of tissue cysts in the muscles and

brain. These are transmitted only if ingested by predation or

scavenging or if passed vertically across the placenta from mother

to foetus.

The localisation of T. gondii cysts in the forebrain of rats and mice

together with the immune reaction to the cysts is related to altered

behaviour, in particular the attenuation of predator odour aversion

and anxiety. This facilitates ingestion of the intermediate host by the

cat definitive host and perpetuation of the life cycle17.

Humans become infected with T. gondii mainly by ingesting un-

cooked meat containing viable tissue cysts or by ingesting food or

water contaminated with oocysts from the faeces of infected cats.

Transplacental transmission may occur in women during their first

trimester of pregnancy and this infection is then passed to the

developingembryovia theplacenta. This frequently results in severe

brain and eye lesions in the newborn or death of the developing

embryo. Research results from the past 15 years have shown that

T. gondii infection is associated with several neuropsychiatric

diseases and behavioural changes in humans as well animals18.

Although the mechanisms are unknown a growing body of data

indicates that they are complex, comprising humoral, immune,

neurotransmitter, epigenetic, genetic, and structural effects19.

Echinococcosis (hydatid disease)

Australia has, on average, >80 new cases of human hydatidosis per

annum caused by the larval stage of the cestode, Echinococcus

granulosus20. The parasite was introduced into Australia with

domestic livestock and dogs. However, a cycle in wildlife is main-

tained through a predator/prey interaction between dingoes, wild

dogs and less importantly foxes and kangaroos and wallabies, less

importantly feral pigs (Sus scrofa)21,22. The establishment of a

dingo/wild dog-macropod cycle, which effectively maintains para-

site transmission, acts as a spill-back reservoir of infection for sheep

and cattle23. This is a major problem for control strategies focussed

onhumaneducation andhusbandrypractices to break thedomestic

‘dog-sheep’ cycle. In contrast to the situation in livestock, hydatid

cysts occur primarily in the lungs rather than the liver of marsupials.

Infectionoccurspredominantly in theeasternStatesbut theparasite

has established recently in wildlife in water catchment and forestry

areas outside Perth21. Western grey kangaroos and feral pigs act as

intermediate hosts. This focus of transmission may have been

initiated through E. granulosus-infected domestic pig hunting dogs

Table 1. Potential zoonotic tissue parasite infections in Australia.

Disease name Organism Type Source Reference

Leishmaniasis Leishmania sp. Trypanosomatid Forcipomyia spp. 26,27

Protozoan

Sparganosis Spirometra erinaceieuropaei Cestode (larval, plerocercoid) Eating raw/undercooked meat 28

Pentastomiasis Armillifer spp. Modified brachiuran Eating undercooked snake/mammal flesh, drinkingcontaminated water

29,30,31

Linguatula spp. Crustacean

Under theMicroscope


from the eastern states. Transmission appears to be perpetuated by

dogs of local pig hunters infected through being fed the offal of

locally shot kangaroos21. Hydatid disease is also an important

conservation issue, especially for small endangered species and

populations of Macropodidae by severely reducing effective lung

volume24,25. Such reductions impact the fitness of the animals

enhancing susceptibility to predation, thus ensuring perpetuation

of the cycle.

Several potential zoonotic tissue parasite infections in Australia are

listed in Table 1.

References1. Mackerras, M.J. and Sandars, D.F. (1955) The life history of the rat lung-worm,

Angiostrongylus cantonensis (Chen) (Nematoda: Metastrongylidae). Aust.

J. Zool. 3, 1–21. doi:10.1071/ZO9550001

2. Spratt, D.M. (2005a) Australian ecosystems, capricious food chains and parasitic

consequences for people. Int. J. Parasitol.35, 717–724. doi:10.1016/j.ijpara.2005.

01.014

3. Spratt, D.M. (2005b) Neuroangiostrongyliasis: disease in wildlife and humans.

Microbiol. Aust. 26, 63–64.

4. Ma, G. (2013) Tawny frogmouths and brushtail possums as sentinels for Angios-

trongylus cantonensis, the rat lungworm. Vet. Parasitol. 192, 158–165.

doi:10.1016/j.vetpar.2012.11.009

5. Prociv, P. and Tiernan, J.R. (1987) Eosinophilic meningoencephalitis with per-

manent sequelae. Med. J. Aust. 147, 294–295.

6. Prociv, P. et al. (2000) Neuro-angiostrongyliasis: unresolved issues. Int.

J. Parasitol. 30, 1295–1303. doi:10.1016/S0020-7519(00)00133-8

7. Morton, N.J. et al. (2013) Severe haemorrhagic meningoencephalitis due to

Angiostrongylus cantonensis among young children in Sydney, Australia. Clin.

Infect. Dis. 57, 1158–1161. doi:10.1093/cid/cit444

8. Senanayake, S.N. (2003) First case of human angiostrongyliasis acquired in

Sydney. Med. J. Aust. 179, 430–431.

9. Blair, N.F. et al. (2013) Angiostrongylus meningoencephalitis: survival from

minimally conscious state to rehabilitation. Med. J. Aust. 198, 440–442.

doi:10.5694/mja12.11085

10. Spratt, D.M. et al. (1999) Haycocknema perplexum n.g., n. sp. (Nematoda:

Robertdollfusidae): an intramyofibre parasite in man. Syst. Parasitol. 43,

123–131. doi:10.1023/A:1006158218854

11. Dennett, X. et al. (1998) Polymyositis caused by a new nematode. Med. J. Aust.

168, 226–227.

12. Basuroy, R. et al. (2008) Parasitic myositis in tropical Australia. Med. J. Aust. 188,

254–256.

13. McKelvie, P. et al. (2013) A further patient with parasitic myositis due to Hay-

cocknemaperplexum, a rare entity. J. Clin.Neurosci.20, 1019–1022.doi:10.1016/

j.jocn.2012.08.009

14. Anderson, R.C. et al. (1998) Halicephalobus gingivalis (Stefanski, 1954) from a

fatal infection of a horse in Ontario, Canada with comments on the validity of

H. deletrix and a review of the genus. Parasite 5, 255–261. doi:10.1051/parasite/

1998053255

15. Lim, C.K. et al. (2015) First human case of fatal Halicephalobus gingivalis

meningoencephalitis in Australia. J. Clin. Microbiol. 53, 1768–1774. doi:10.1128/

JCM.00032-15

16. Blunden, A.S. et al. (1987) Halicephalobus deletrix infection in a horse. Equine

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clinids/9.6.1087

Biography

Dave Spratt is an Honorary Fellow at the Australian National

Wildlife Collection, National Research Collections Australia, CSIRO,

in Canberra. Themajor themes of Dr Spratt’s research are the study

and understanding of the diseases of wildlife including disease

ecology, parasite taxonomy, helminth biodiversity and zoonoses.

Research topics of interest includemetastrongyloid, filarioid, trichi-

nelloid and muspiceoid nematodes, pentastomes, and small mam-

mal succession and recolonisation of their helminth communities

following wildfire.

Under theMicroscope


Assessing enteric helminths in refugees, asylumseekers and new migrants

Sarah HaniehA,D,*, Norbert RyanB,* and Beverley-Ann BiggsB,C

AVictorian Infectious Diseases Reference Laboratory, Peter Doherty Institute for Immunity and Infection, Parkville, Vic. 3050, Australia

BDepartment of Medicine, The University of Melbourne, Peter Doherty Institute for Immunity and Infection, Parkville, Vic. 3050, Australia

CThe Victorian Infectious Diseases Service, Royal Melbourne Hospital, Parkville, Vic. 3050, Australia

DCorresponding author. Department ofMedicine, The University of Melbourne, Doherty Institute, Parkville, Vic. 3050, Australia, Tel: +61 3 8344 3257,Fax: +61 3 9347 1863, Email: [email protected]

Currently there are 59.5million people forcibly displaced

worldwide as a result of conflict, human rights violations,

generalised violence or persecution. Of these, 19.5million

are refugees and 1.8million are asylum seekers. Each year

Australia accepts 13750 refugees through the offshore

Humanitarian program, and in 2016 that number will

almost double with the addition of 12000 refugees from

Syria and Iraq. Many refugees have complex medical needs

and have reached Australia after a difficult journey, often

involving time in refugee camps and exposure to traumatic

events including physical hardship and illness. Refugees

often come from parts of the world where parasitic and

tropical infectious diseases are prevalent and untreated.

This article provides a review of enteric helminth infections

in refugees, including asylum seekers and those from a

refugee-like background.

Parasitic infections in refugees and new migrants reflect the under-

lying epidemiology of parasites in areas where refugees may have

been exposed, including the country of origin, migration journey to

Australia, andplaceof detention. Factors such aspoverty, disruption

of basic services, poor sanitation/hygiene (e.g. quality of drinking

water, access to running water, access to footwear) and insufficient

access to adequate health care and treatment may significantly

increase the risk of exposure to intestinal parasitic disease in the

refugee population1. Other practices such as the use of night-soil as

fertiliser, dietary habits, and past occupational exposures are likely

to play an important role in increasing the burden of disease. Soil

transmitted helminth (STH) infections are very common in those

living in resource constrained settings1. Treatment of refugees and

asylum seekers is often empirical, in refugee camps, before depar-

ture as part of the pre-departure health check, or after arrival in

Australia. More serious infections, such as strongyloidiasis, schisto-

somiasis, opisthorchus and Taenia solium, require diagnosis

and specific treatment. Table 1 summarises the findings of recent

prevalence studies of strongyloidisis and schistosomiasis in refugee

groups from Australia and overseas.

Strongyloidiasis

The highest prevalence of Strongyloides stercoralis infection

occurs in refugees from Africa and South-East Asia2. Of those

arriving in the last decade, the Burmese groups (e.g. Karen, Chin)

have the highest prevalence (26.0%). Earlier data show an even

*Joint first authors.

Under theMicroscope


higher prevalence in Lao and Cambodian refugees3–5. Infections

may persist for decades after leaving an endemic area due to a

continual cycle of auto-infection. Although many patients are

asymptomatic or have minimal clinical symptoms, they remain at

risk for subsequent hyperinfection if immunosuppressed6. Stron-

gyloides antibodies decline after effective treatment7.

Schistosomiasis

The highest prevalence of schistosomiasis is found in Africa, ac-

counting for an estimated 95% of global cases. Species involved are

S. mansoni, S. haematobium and S. intercalatum. S. japonicum is

present along the Yangtze River in China and the Philippines.

S. mekongi is found in the Mekong river valley. Schistosoma

infection may also be encountered in other areas such as parts of

Indonesia, the Caribbean, the Arabian Peninsula, Madagascar, the

Middle east and Turkey8. Prevalence of schistosomiasis infection in

refugees in Australia, as determined by serology, has been shown

to range from 5.4% in Burmese9 to 37% in Africans10.

Schistosoma antibodies are thought to persist in those from en-

demic areas despite prior treatment. A long-term study of schisto-

somiasis serology post-treatment showed an immediate increase in

titre and thena fourfolddecline inmost travellers after 6–12months.

Table 1. Prevalence of strongyloidiasis and schistosomiasis in refugee and asylum seeker groups.

Reference Sample size Country/regionof origin

Country ofsettlement

Prevalence ofSchistosomiasis

Prevalence ofStrongyloides

International studies

21 700 Latin America, Sub-saharan Africa

Spain 5.9% 56.1%

22 1063 Europe, EasternMediterranean, Africa

Canada 15% 3%

23 208 Brazil USA 27.7% 5.8%

24 350 Africa (46%), Asia(28.6%)

Canada N/A 4.6%

16 1376 Middle East, Africa,Asia

USA 0.8% (Africa) 2.0% (SE Asia)2.5% (Africa)

20 176 Southeast Asia (59%),Africa (27%), MiddleEast (14%)

USA 8% 24%

Australian studies

21 1136 Burma (Karenrefugees)

Australia 7% 20.8%

21 187 Asia Australia 17% 5.7%

27 182 Africa Australia N/A N/A

7 234 Cambodia Australia N/A 35%

9 156 Burma Australia 5.4% 26%

10 239 Africa Australia 37% –

28 258 Sudan, Liberia Australia 12% 9%

3 361 Cambodia, East Africa Australia 11% East African 2% East African42% Cambodian

29 135 East Africa Australia 2% 11%

5 95 Laos Australia N/A 23%

N/A, not available.

Under theMicroscope


However, for immigrants, serology remained elevated even

three years after effective treatment in a proportion of patients11.

Soil-transmitted helminths (STH)

Many refugees will have received empirical albendazole as part of

the predeparture medical assessment conducted by the Interna-

tional Organisation of Migration on behalf of the Australian

Government. This has significantly altered the prevalence and

patternsof intestinal helminths in refugees12.However, albendazole

has limited effectiveness against Trichuris trichiura, and is not an

effective treatment for some less common helminths found in

refugees and asylum seekers (see below).

Less common helminth infections

Themajority of Asian refugees currently entering Australia are from

Myanmar, from camps on the Thai border. Within Thailand, espe-

cially the north east, Opisthorchis viverrini is highly prevalent.

Infection results from consumption of raw, uncooked or fermented

fish containing metacerciae. Long-term infection may cause cho-

langitis, obstructive jaundice, cholecystitis, periductal fibrosis and

bile duct cancer, contributing to a liver cancer rate in excess of

70 per 100 000 in NE Thailand. There is a paucity of data on faecal

microscopy findings for refugees from Myanmar. However, the

presence of vector cyprinoid fish and substantial wetlands suggests

that infection with Opisthorchis viverrini is also likely.

Other serious infections in refugees from Asia include Taenia

solium (pork tapeworm) with the potential risk of cysticercosis for

both patient and household members. In Thailand faecal tests

for helminth eggs have revealed a prevalence of Taenia spp of

2.3–3.7% in communities with high migrant populations along the

Thai border. In the remote western border area of Kanchanaburi

three species of tapeworm T. saginata, T. solium and T. asiatica

co-exist in the human population13.

Considerations in the Syrian refugee population

In September 2015, it was announced that 12 000 Syrian and Iraqi

refugees would be accepted to Australia as part of theHumanitarian

Program during 2016–17. The recent prevalence of enteric parasite

infections is not well documented in Syria. However, schistosomi-

asis is considered to have low endemnicity in Iraq (0.1%) and

Syria (<10%prevalence in 2010)14. There is no information available

on the prevalence of S. stercoralis in Syria; however, a hospital-

based survey in Iraq reported a prevalence of 24.2%15. Chang et al.

reported a low prevalence of other parasitic infections in refugees

from Iran and Iraq16.

Diagnosis

In Australia, the number of faecal microscopy tests performed has

fallen in some States (e.g. NSW), with more emphasis being placed

on empirical treatment of STH and the use of serology for the

diagnosis of S. stercoralis or Schistosoma spp. Current recom-

mended diagnostic tests for enteric parasites are shown in Table 2.

Challenges and limitations of testing for

schistosoma and strongyloides infections

in a refugee population

Serology may overestimate the prevalence of disease due to cross-

reactivity with other nematode infections and there is difficulty

distinguishing recent from past (and cured) infections. Serological

titres (OD values) in the equivocal and low positive ranges are

difficult to interpret. Follow-up serology should preferably be done

in the same laboratory and in parallel with previous specimens

where available. The interpretation of Schistosoma serology in

Table 2. Recommended diagnostic tests for enteric parasites.

Enteric parasite Diagnosis

Strongyloides stercoralis Positive serologyStool microscopy should be performed to rule out other enteric infections

Schistosoma spp. Positive serologyStool/urine microscopy for ova should be performed in those with positive serology

Hookworm (Necator americanus and Ancylostoma duodenale) Microscopic finding of ova in faecal specimensConcentration methods are necessary to detect light infections

Ascaris lumbricoides Microscopic identification of ova in faecal specimens

Trichuris trichiura Microscopic identification of ova in faecal specimens

Taenia spp. Microscopic identification of ova and proglottids in faecal specimens

Under theMicroscope


Australia presents several challenges as test methodology varies

between laboratories. VIDRL in Melbourne, use the Fumouze

indirect haemagglutination test (IHA). This is based on an antigen

derived from adult worms of S. mansoni. Estimates in one com-

parison study show that the sensitivity of this test is 76.2% for

Schistosomamansoni, and slightly lower for S. haematobium, with

a specificity of 99%17. However, in NSW at ICPMR, an ‘in house’

ELISA assay is the preferred assay used, based on S. mansoni egg

antigen. Two commercial assays based on the use of a similar

antigen showed 71.4–85.7% sensitivity but reduced specificity of

76.9–88.4%. Both ELISA assays showed cross-reactivity with ces-

tode, nematode and trematode infections17. The sensitivity of these

assays for other species of Schistosoma, such as S. haematobium,

S. intercalatum, S. mekongi and S. japonicum, is not specified;

however, it is likely to be reduced.

The sensitivity and specificity of Strongyloides stercoralis serology

is reported to be up to 94.6% and 99.6% respectively, depending

on the assay used18. However, as there is no gold standard test for

comparison, these are estimations only. Serological titres decline

with effective treatment over a 12 month period7,18.

Persistence of parasitic infections in refugee

populations

Several studies have demonstrated that serious intestinal parasitic

infectionsmay persist formany years after arrival in Australia. A 2002

study of Laotian refugees who had arrived in Australia during

1974–91 showed that strongyloides serology was positive in 24%

and 3 carried Opisthorchis, compared to respective prevalences

of 19.2% and 41%, on initial screening by faecal microscopy4. As

liver flukes survive for approximately 7–10 years, the three cases of

Opisthorchis identified may well represent reinfection on subse-

quent visits to Laos.

In a second study of East African refugees, who arrived in Australia

in the late 90s, screening for strongyloides and schistosoma

serology was positive in 11% and 15%, respectively, of patients

some 16 years later in 20063. In a further aspect of the same study,

42% of 234 Cambodians who had arrived in the late 80s still tested

positive for strongyloides serology3. This compared with 7.8% of

Cambodians who were found to be positive for strongyloides by

microscopy at initial health screening4.

Post arrival health assessment for refugees

and asylum seekers

Community Health Centres and GP services in suburbs with high

rates of migrant and refugee settlement are now responsible for

much of the post-arrival refugee health screening, with support

from specialist Refugee Health Services. Guidelines for post-arrival

assessment for people of refugee-like background were published

by the Australian Society for Infectious Diseases in 2009 and are

currently being revised19. If possible, a full health assessment of new

arrivals is ideally conducted within onemonth of arrival in Australia,

including serology for Strongyloides for all, and Schistosoma in

those who have lived or travelled through an endemic area. For

many of the clinics in suburbs with high migrant populations,

the special pathology requirements are met by private pathology

providers.

Summary

Intestinal helminth infections in refugees are common and should

remain a high priority for health workers. These populations often

have specific needs that should be considered in diagnosis and

managementof these infections.Burdenofdisease is likely to reflect

the country of origin, journey of migration to Australia, pre-depar-

ture treatment and place of detention. Other socio-economic and

cultural factors are also likely to play a significant role in exposure

risk. Helminth infections may be chronic and persist in humans for

more than four decades resulting in seriousmorbidity andmortality

and highlighting the need for early diagnosis.

Acknowledgements

We thank Ms Christalla Hajisava for technical assistance.

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Biographies

Dr Sarah Hanieh is a paediatric infectious diseases physician and

NHMRC Early Career Research Fellow in the Immigrant and Inter-

national Health Group, at the Peter Doherty Institute for Infection

and Immunity, The University of Melbourne. She has interests in

child nutrition, infectious diseases and refugee health. Her current

research aims to understand the nutritional and infectious causes of

impaired child growth and development in resource poor settings.

Norbert Ryan is a Senior Scientist in the Bacteriology Laboratory at

Victorian Infectious Diseases Reference Laboratory, Doherty Insti-

tute, Melbourne. Areas of interest include parasitology, diagnosis of

Legionnaires’diseaseandsexually transmitted infections.Theuseof

staining methods for diagnosis of protozoal infections including

microsporidia has been a specialty field. He was involved in proces-

sing of specimens during the systematic health screening of refugee

and family reunion migrant groups conducted by the Victorian

Department of health during the late 80s-early 90s.

Professor Beverley-Ann Biggs heads the International and Im-

migrant Health Group in the Department of Medicine, The Univer-

sity of Melbourne and is an Infectious Diseases Physician in the

Victorian Infectious Diseases Service at the Royal Melbourne

Hospital. She has a special interest in parasitic and other infectious

diseases in refugees and immigrants living in Australia, and has

published extensively in this area.

Under theMicroscope


Free living amoebae and human disease

Evan Bursle

Sullivan Nicolaides PathologyWhitmore StreetTaringa, Qld 4068, AustraliaEmail: [email protected]

Jennifer Robson

Sullivan Nicolaides PathologyWhitmore StreetTaringa, Qld 4068, AustraliaEmail: [email protected]

Pathogenic FLA are ubiquitous protozoans and despite fre-

quent human contact remain a rare cause of often devastat-

ing infectionwith poor prognosis. Given changes in climate,

human encroachment into the environment, increasing im-

munosuppression, and improving diagnostic capacity, it is

likely we will see increased cases in the future. Early diag-

nosis is challenging but crucial to achieving a favourable

outcome. It is best facilitated by improved awareness of

FLA disease, appropriate clinical suspicion and early diag-

nostic testing.

Free living amoebae (FLA) are a cosmopolitan group of protozoan

organisms that do not require a host to survive. Despite their

ubiquitous nature, these organisms are uncommon human patho-

gens.However, four genera contain species known to cause invasive

disease in humans: Acanthamoeba, Naegleria, Balamuthia and

Sappinia. Acanthamoeba infectionsmaypresent as granulomatous

amoebic encephalitis (GAE), disseminated disease (e.g. cutaneous,

sinus or pulmonary infection) or keratitis, with Balamuthia man-

drillaris causing similar cutaneous infections and GAE. Naegleria

fowleri is responsible for the rapidly progressive primary amoebic

meningoencephalitis (PAM). In addition, a single case of human

central nervous system (CNS) infection with Sappinia pedata has

been reported1, along with isolated cases of corneal infection with

Vahlkampfia spp.,Hartmannella spp. and Paravahlkamfia spp.2.

This review aims to provide an overview of human disease caused

by the three most common genera involved, Acanthamoeba,

Naegleria and Balamuthia, including their laboratory diagnosis.

Epidemiology

Pathogenic FLAare foundworldwideand serological studies suggest

humanexposure is common3–5. Three cases ofAcanthamoebaGAE

have been described in Australia6–8. Acanthamoeba sp tolerate a

wide rangeof temperature, pH andosmolarity9 andmay be found in

air, soil and water samples. They are one of the most commonly

isolated FLA in the environment10, and themost common in human

infection2. The strongest risk factor for GAE or disseminated infec-

tion is immunodeficiency, while keratitis most commonly affects

immunocompetent contact lenswearerswhooftenhave ahistory of

poor lens hygiene. Exposure is thought to occur by inhalation,

mucosal contact or direct inoculation10. Acanthamoeba species

were previously classified based on morphology, but are now

grouped into 17 genotypes based on 18S rRNA sequencing, with

the majority of pathogenic species belonging to the T4 genotype9,11.

Naegleria fowleri may be found in rivers, lakes and soil but does

not survive in sea water. As thermophiles, their presence in fresh

water is related to temperature and they may even be recovered

from thermally pollutedwaters at high latitudes12. Human exposure

occurs through contact with intact or disrupted nasal mucosa,

commonly through recreational or nasal ablution practices13 and

contaminated drinking water has been implicated as a source of

infection in some cases14. Australia has featured prominently in

the history of Naegleria fowleri infection, with cases described in

Queensland, New South Wales and Western Australia following the

first description of the disease in 1965 by two South Australian

pathologists, Fowler and Carter15,16. This discovery was related to

an outbreak of 20 cases, attributed to a contaminated overland

water pipeline which reached optimal temperatures for Naegleria

proliferation during the summer months. Cases continued from

1947 to 1972 when public health measures including adequate

chlorination were applied17. Several cases inWestern Australia have

also been associated with overland water pipelines, and Australian

drinking water guidelines suggest a Naegleria monitoring and

response protocol for water supplies that seasonally exceed 308C,

or 258C continually18. Today, despite significant advancement in

Under theMicroscope


knowledge regarding risk reduction measures, the incidence of

infection with Naegleria fowleri appears to be increasing world-

wide, with factors such as global warming, substandard water

management and sanitation, and changing recreational practices

likely involved13.

Balmuthia mandrillaris is most commonly found in soil, though

it may also be recovered from water samples4. Infection tends to

occur in immunocompetent individuals, most commonly children,

probably through inhalation or nasal or cutaneous inoculation.

Transmission via organ transplantation has also been described19.

Six human cases have been described in Australia20–24 (Western

Australia, Tasmania, Victoria and Queensland), with another being

described in a Victorian dog25.

Clinical manifestations

Acanthamoeba sp.

The predominant clinicalmanifestations of Acanthamoeba infection

are disseminated disease (e.g. cutaneous, nasopharyngeal or pulmo-

nary infection), GAE and keratitis. With the exception of rare case

reports, GAE and disseminated disease occur in the immunocom-

promised or debilitated. The incubation period is unknown, but

thought to be weeks to months in duration. Cutaneous infection

usually begins as fibronodular lesions, which progress to non-healing

ulcerated lesions over time10. While GAE may be fatal within days of

symptom onset, it generally assumes a more chronic course, with

slow progression over weeks to months. Clinical features of GAE

are myriad and include fever, symptoms of meningism, personality

and mental status change and, later, focal neurological deficits,

coma and death. Imaging of the brain may show single or multiple

space occupying lesions which can be ring enhancing.

Acanthamoeba keratitis is usually a disease of immunocompetent

patients, with the strongest risk factor being contact lens use and

poor lens hygiene. The disease is usually unilateral, with symptoms

including lacrimation, pain, photophobia and foreign body sensa-

tion. Signs include a typical corneal ring infiltrate, stromal infiltrates,

epitheliopathy and hypopyon26. In the absence of effective therapy,

it may progress to corneal perforation and loss of vision. The clinical

diagnosis of AK can be difficult, as lesions may resemble bacterial

or fungal disease, or the dendritic ulcer of HSV infection26. Further-

more the clinical course may be characterised by periods of tem-

porary remission, leading to false impressions of response to

antibacterial or viral agents2.

Naegleria fowleri

Naegleria fowleri causes primary amoebic meningoencephalitis.

Symptoms generally occur 2–5 days after exposure and may begin

with changes in taste or smell, followed by fever, nausea, vomiting,

photophobia and headache. The disease is fulminant, with rapid

progression to coma and death.

Balamuthia mandrillaris

Balamuthia mandrillaris causes GAE in immunocompromised

and immunocompetent individuals. The onset of meningoenceph-

alitis is often subacute or chronic, with symptoms developing over a

period of 2 weeks to 2 years27. It also has the propensity to cause

cutaneous lesions that may precede CNS involvement and are

similar in appearance to those of Acanthamoeba sp. These lesions

appear as poorly defined plaques and may be single or with

bordering satellite lesions4. They often involve the central face and

appear to be more common in South America4. Cutaneous disease

generally progresses to CNS involvement; however, it may resolve

with therapy4.

Laboratory diagnosis

Diagnosis of FLA infection, particularly systemic disease, is chal-

lenging: it may masquerade as bacterial or viral infection, exposure

events may not be apparent and specialised diagnostic testing

availability is limited. Unfortunately as a result, many CNS infections

are diagnosed post-mortem. Successful early diagnosis depends on

appropriate clinical suspicion and collection of suitable diagnostic

material, usually tissue or CSF.

Microscopy

CSF samples in cases of PAM appear purulent, with no bacteria

evident on gram stain, a polymorphonuclear pleocytosis, elevated

protein and decreased glucose. Wet preparations may show motile

Naegleria fowleri trophozoites (Figure 1a), as there are usually

large numbers of organisms in theCSF. Thesefindings contrast with

those of GAE caused by Acanthamoeba or Balamuthia. While CSF

from cases of GAE also demonstrates elevated protein and lowered

glucose, these changes aremoremodest, andamononuclear, rather

than polymorphonuclear inflammatory response is seen. Further-

more,AcanthamoebaorBalamuthia trophozoites are not typically

seen in CSF preparations.

Trophozoites from all pathogenic FLA species can be difficult to

differentiate from host inflammatory cells, especially in stained

tissue sections. Thenuclear characteristics of amoeba canbehelpful

in differentiating these parasites from host cells, with Naegleria

fowleri possessing a nucleus with a large, round, central nucleolus

and Acanthamoeba and Balamuthia (Figure 1b) a rounded

nucleus with large, central nucleolus forming a halo. Polyclonal

andmonoclonal antibodies, with a secondary detecting fluorescent

anti-IgG antibody (such as FITC), may be used to identify and

Under theMicroscope


differentiate each of these amoebae in tissue specimens28 (as can

molecular methods), though availability is limited to the CDC,

Atlanta, USA.

In Acanthamoeba keratitis, a diagnosis may be made by demon-

strating trophozoites and/or cysts in corneal samples. It is possible

to directly identify Acanthamoeba trophozoites within the cornea

using confocalmicroscopy and in experienced hands this technique

is sensitive and specific29. Trophozoites and cysts may be revealed

by staining the smear with H&E or Giemsa, while cysts are also

readily identifiedusingPASandfluorescent stains, suchascalcofluor

white and acridine orange30. On occasion, non-specific fluores-

cence or binding to fungi in mixed infections can lead to diagnostic

errors31, especially when used by inexperienced microscopists.

Culture

Samples intended for amoebic culture should be kept at room

temperature and processed as quickly as possible. Freezing should

be avoided, particularly for samples where Naegleria is suspected

(the cyst stage is more fragile), as this compromises organism

viability. Naegleria fowleri and Acanthamoeba sp. (Figure 1c) can

be readily cultured using non-nutrient media containing live or

killed non-mucoid bacteria (usually E. coli or Enterobacter sp) as a

food source32. Acanthamoeba and Naegleria will cover the agar

surface in 1–2 days when incubated at 378C and their presence can

be confirmed by examination of the plate with a plate microscope,

or by performing microscopy of a wet mount from the agar plate.

Balamuthia do not appear to use bacteria as a food source and

therefore cannot be cultivated in the same fashion32. They may be

successfully cultured using axenic and tissue culture methods32,

however with generation times of around 25 h, culture is a lengthy

process and not part of routine diagnostic testing.

Nucleic acid testing

The use of molecular testing to diagnose and confirm infections

with FLA has transformed diagnostics in this area. It allows more

rapid diagnosis, with greater sensitivity than other methods and

reduces the requirement for specialist, experienced staff to discern

subtle microscopic features. Most molecular assays use ribosomal

genes, such as the 18S rRNA or ITS repeat regions, as targets for

PCR. However, as requests are infrequent, these tests are generally

only offered by reference or research laboratories. Of particular

note is a multiplex PCR, described by Qvarnstrom et al.33. This

assay uses 18S rRNA primer/probe sets to accurately identify

Acanthamoeba to the genus level and Naegleria fowleri and

Balamuthia mandrillaris species. The sensitivity is reported at

one amoeba per sample. Sullivan Nicolaides Pathology instituted

(a) (b) (c)

Figure 1. (a) Naegleria trophozoite in CSF wet prep (x400). (b) Balamuthia trophozoite in brain tissue (haematoxylin and eosin). (c) Acanthamoebacysts in culture, wet prep (x400).

Table 1. Multiplex Free Living Amoebae PCR33 at Sullivan Nicolaides Pathology, November 2011 to July 2015A.

Organism Eye CNS Skin Sinus Total

Acanthamoeba spp 5 1 81

Balamuthiamandrillaris

1 63

Naegleria fowleri 2 63

Total tests 52 24 4 1 81

ADuplicates excluded (same patient and site, within 2 months). CNS, central nervous system.

Under theMicroscope


this test in 2010 and our experience is summarised in

Table 1. As a significant number of ocular specimens have been

tested against all three targets, a large number of negatives for

Naegleria fowleri and Balamuthiamandrillaris are expected. It is

alsonotable thatwehave recently reported aprobable false negative

Acanthamoeba result. The referred CSF specimen tested negative

at our laboratory, but positive at the CDC (using the same

multiplex PCR). It is possible the small volume of CSF received at

our laboratory contributed to this result (only 100mL was received,

where 200mL is the standard volume for extraction).

Treatment and prognosis

Primary amoebic meningitis

There have been few survivors of primary amoebic meningitis

and the factors that have resulted in successful therapy are poorly

defined, though early diagnosis and institution of therapy appears

critical. The treatment of choice is the antifungal amphotericin B,

which is often used both intravenously and intrathecally. Other

agents used in survivors include miltefosine, sulfisoxazole, flucon-

azole, miconazole and rifampicin.

Granulomatous amoebic encephalitis

Acanthamoeba

Suboptimal efficacy of antimicrobial agents, the high morbidity of

patients affected and tendency to late diagnosis contribute to a

poor prognosis in Acanthamoeba GAE, with a mortality rate of

>90%34. In the few reported survivors of GAE or cutaneous

infection, most have received combination therapy. The agents

used have included trimethoprim-sulfamethoxazole, flucytosine,

sulfadiazine, penicillin G, chloramphenicol, pentamidine, flucona-

zole and itraconazole. More recently, it appears that the inclusion

of miltefosine in combination regimens may result in improved

survival35.

Balamuthia

Early recognition of cutaneous disease is critical to allow early

therapy and prevent progression to CNS infection. Unfortunately,

the overall prognosis in Balamuthia CNS infection is extremely

poor. However, there are several case reports of survival in the

literature36, including an Australian case24. These cases have re-

ceived varied combination therapy regimens, with agents including

pentamidine, flucytosine, fluconazole, macrolides, sulfadiazine,

miltefosine, thioridazine, amphotericin B, albendazole and trimeth-

oprim-sulfamethoxazole. Miltefosine, in particular, has demonstrat-

ed amoebicidal activity in vitro37 and its inclusion in combination

regimens may offer a survival advantage35.

Acanthamoeba keratitis

Like disseminated disease, success in treatment is dependent on

early diagnosis and institution of therapy. Fortunately, success

rates in treating Acanthamoebic keratitis are more promising, with

cure rates in the literature generally greater than 75–85%38,39.

Topical chlorhexidine and polyhexamethylenebiguanide (PHMB)

are effective against trophozoites and cysts and form the mainstay

of therapy. These are usually used in combination with diamidine,

propamidine or hexamidine, though other agents including keto-

conazole, itraconazole, voriconazole and topical imidazoles have

been used. Surgical intervention, including enucleation, is some-

times required in severe cases40.

References1. Qvarnstrom, Y. et al. (2009) Molecular confirmation of Sappinia pedata as a

causative agent of amoebic encephalitis. J. Infect. Dis. 199, 1139–1142.

doi:10.1086/597473

2. Mandell, G.L. et al. (2015) Mandell, Douglas, and Bennett’s principles and

practice of infectious diseases. Philadelphia, PA: Churchill Livingstone/Elsevier.

3. Chappell, C.L. et al. (2001) Standardized method of measuring Acanthamoeba

antibodies in sera from healthy human subjects. Clin. Diagn. Lab. Immunol. 8,

724–730.

4. Bravo, F.G. and Seas, C. (2012) Balamuthia mandrillaris amoebic encephalitis:

an emerging parasitic infection. Curr. Infect. Dis. Rep. 14, 391–396. doi:10.1007/

s11908-012-0266-4

5. Marciano-Cabral, F. et al. (1987) Specificity of antibodies from human sera for

Naegleria species. J. Clin. Microbiol. 25, 692–697.

6. Carter, R.F. et al. (1981) A fatal case of meningoencephalitis due to a free-living

amoeba of uncertain identity–probably Acanthamoeba Sp. Pathology 13, 51–68.

doi:10.3109/00313028109086829

7. Harwood, C.R. et al. (1988) Isolation of Acanthamoeba from a cerebral abscess.

Med. J. Aust. 148, 47.

8. Azzam, R. et al. (2015) Acanthamoeba encephalitis: isolation of genotype T1

in mycobacterial liquid culture medium. J. Clin. Microbiol. 53, 735–739.

doi:10.1128/JCM.02887-14

9. Trabelsi, H. et al. (2012) Pathogenic free-living amoebae: epidemiology

and clinical review. Pathol. Biol. (Paris) 60, 399–405. doi:10.1016/j.patbio.

2012.03.002

10. Marciano-Cabral, F. and Cabral, G. (2003) Acanthamoeba spp. as agents of

disease in humans. Clin. Microbiol. Rev. 16, 273–307. doi:10.1128/CMR.16.

2.273-307.2003

11. Siddiqui, R. and Khan, N.A. (2012) Biology and pathogenesis of Acanthamoeba.

Parasit. Vectors 5, 6. doi:10.1186/1756-3305-5-6

12. Kemble, S.K. et al. (2012) Fatal Naegleria fowleri infection acquired in

Minnesota: possible expanded range of a deadly thermophilic organism. Clin.

Infect. Dis. 54, 805–809. doi:10.1093/cid/cir961

13. Siddiqui, R. and Khan, N.A. (2014) Primary amoebic meningoencephalitis

caused by Naegleria fowleri: an old enemy presenting new challenges. PLoS


14. Cooter, R. (2002) The history of the discovery of primary amoebic meningoen-

cephalitis. Aust. Fam. Physician 31, 399–400.

15. Fowler, M. and Carter, R.F. (1965) Acute pyogenic meningitis probably due to

Acanthamoeba sp.: a preliminary report. BMJ 2, 734–742. doi:10.1136/bmj.2.

5464.734-a

16. Carter, R.F. (1968) Primary amoebic meningo-encephalitis: clinical, pathological

and epidemiological features of six fatal cases. J. Pathol. Bacteriol. 96, 1–25.

doi:10.1002/path.1700960102

Under theMicroscope


17. Dorsch, M.M. et al. (1983) The epidemiology and control of primary amoebic

meningoencephalitis with particular reference to South Australia. Trans. R. Soc.

Trop. Med. Hyg. 77, 372–377. doi:10.1016/0035-9203(83)90167-0

18. National Health and Medical Research Council Australia (2011) Natural

Resource Management Ministerial C. Australian drinking water guidelines 2011:

national water quality management strategy. Canberra: National Health and

Medical Research Council.

19. Schlessinger, S. et al. (2010) Balamuthia mandrillaris transmitted through

organ transplantation – Mississippi, 2009. Morbidity and Mortality Weekly

Report 59, 1165–1170.

20. Carter, R.F. et al. (1981) A fatal case of meningoencephalitis due to a free-living

amoeba of uncertain identity–probably Acanthamoeba sp. Pathology 13, 51–68.

doi:10.3109/00313028109086829

21. Reed, R.P. et al. (1997) Fatal granulomatous amoebic encephalitis caused by

Balamuthia mandrillaris. Med. J. Aust. 167, 82–84.

22. Hill, C.P. et al. (2011) Balamuthia amebic meningoencephalitis and mycotic

aneurysms in an infant. Pediatr. Neurol. 45, 45–48. doi:10.1016/j.pediatrneurol.

2011.05.003

23. Doyle, J.S. et al. (2011) Balamuthia mandrillaris brain abscess successfully

treated with complete surgical excision and prolonged combination antimicrobial

therapy. J. Neurosurg. 114, 458–462. doi:10.3171/2010.10.JNS10677

24. Moriarty, P. et al. (2014) Balamuthia mandrillaris encephalitis: survival of a

child with severe meningoencephalitis and review of the literature. J. Pediatric

Infect. Dis. Soc. 3, e4–e9. doi:10.1093/jpids/pit033

25. Finnin, P.J. et al. (2007) Multifocal Balamuthia mandrillaris infection in a dog

in Australia. Parasitol. Res. 100, 423–426. doi:10.1007/s00436-006-0302-0

26. Patel, D.V. and McGhee, C.N. (2009) Acanthamoeba keratitis: a comprehensive

photographic reference of common and uncommon signs. Clin. Experiment.

Ophthalmol. 37, 232–238. doi:10.1111/j.1442-9071.2008.01913.x

27. Visvesvara, G.S. et al. (2007) Pathogenic and opportunistic free-living

amoebae: Acanthamoeba spp., Balamuthia mandrillaris, Naegleria fowleri,

and Sappinia diploidea. FEMS Immunol. Med. Microbiol. 50, 1–26. doi:10.1111/

j.1574-695X.2007.00232.x

28. da Rocha-Azevedo, B, Tanowitz, HB and Marciano-Cabral, F (2009) Diagnosis

of infections caused by pathogenic free-living amoebae. Interdisciplinary

perspectives on infectious diseases 2009, 251406. doi:10.1155/2009/251406

29. Tu, E.Y. et al. (2008) The relative value of confocal microscopy and superficial

corneal scrapings in the diagnosis of Acanthamoeba keratitis. Cornea 27,

764–772. doi:10.1097/ICO.0b013e31816f27bf

30. Hahn, T.-W. et al. (1998) Acridine orange staining for rapid diagnosis of Acantha-

moeba keratitis. Jpn. J. Ophthalmol. 42, 108–114. doi:10.1016/S0021-5155(97)

00127-5

31. Grossniklaus, H.E. et al. (2003) Evaluation of hematoxylin and eosin and special

stains for the detection of Acanthamoeba keratitis in penetrating keratoplasties.

Am. J. Ophthalmol. 136, 520–526. doi:10.1016/S0002-9394(03)00322-2

32. Schuster, F.L. (2002) Cultivation of pathogenic and opportunistic free-living

amebas. Clin. Microbiol. Rev. 15, 342–354. doi:10.1128/CMR.15.3.342-354.2002

33. Qvarnstrom, Y. et al. (2006) Multiplex real-time PCR assay for simultaneous

detection of Acanthamoeba spp., Balamuthia mandrillaris, and Naegleria

fowleri. J. Clin. Microbiol. 44, 3589–3595. doi:10.1128/JCM.00875-06

34. Barratt, J.L.N. et al. (2010) Importance of nonenteric protozoan infections in

immunocompromised people. Clin. Microbiol. Rev. 23, 795–836. doi:10.1128/

CMR.00001-10

35. Cope, J. et al. (2013) Increased patient survival: miltefosine for treatment of

free-living ameba infections caused by Acanthamoeba and Balamuthia

[Poster]. Council of State and Territorial Epidemiologists’ Annual Conference.

Pasadena, California.

36. Deetz, T.R. et al. (2003) Successful treatment of Balamuthia amoebic

encephalitis: presentation of 2 cases. Clin. Infect. Dis. 37, 1304–1312.

doi:10.1086/379020

37. Schuster, F.L. et al. (2006) In-vitro activity of miltefosine and voriconazole on

clinical isolates of free-living amebas: Balamuthia mandrillaris, Acanthamoeba

spp., and Naegleria fowleri. J. Eukaryot. Microbiol. 53, 121–126. doi:10.1111/

j.1550-7408.2005.00082.x

38. Dart, J. K. et al. (2009) Acanthamoeba keratitis: diagnosis and treatment update

2009. Am. J. Ophthalmol. 148, 487–99e2. doi:10.1016/j.ajo.2009.06.009

39. Butler, T.K.H. et al. (2005) Six-year review of Acanthamoeba keratitis in New

South Wales, Australia: 1997–2002. Clin. Experiment. Ophthalmol. 33, 41–46.

doi:10.1111/j.1442-9071.2004.00911.x

40. Vemuganti, G.K. et al. (2005) Granulomatous inflammation in Acanthamoeba

keratitis: an immunohistochemical study of five cases and review of literature.

Indian J. Med. Microbiol. 23, 231–238.

Biographies

Dr Evan Bursle, BSc, MBBS, is a microbiology and infectious

diseases registrar based at Sullivan Nicolaides Pathology. Although

his microbiological tastes are broad and still being refined, he has a

keen interest in parasitology.

Dr Jenny Robson, MBBS (Hons I); FRACP, FRCPA, FACTM, is an

infectious disease physician andmicrobiologist who has worked for

the past 26 years at Sullivan Nicolaides Pathology. She has a broad

range of interests, which includes travel and tropical medicine.

Future issues of Microbiology Australia

May 2016: Education to enhance microbiology graduate employability

Guest Editor: Danilla Grando

September 2016: Diseases of Aquaculture

Guest Editor: Nicky Buller

November 2016: Microbiology of Travel

Guest Editor: Ipek Kurtboke; joint issue with the Society for General Microbiology

March 2017: Bat-associated diseases

Guest Editor: Glenn Marsh

Under theMicroscope


Tapeworm cysts in the brain: can we preventit happening?

Marshall Lightowlers

Facultyof VeterinaryandAgriculturalSciencesThe University of Melbourne250 Princes HighwayWerribee, Vic. 3030, AustraliaEmail: [email protected]

Imagine theconsternation; youareamemberofanorthodox

Jewish family and you and another family member are

diagnosed with larvae of a pork tapeworm in your brain.

You have recurrent seizures as a result. Ridiculous? Not

for members of a Jewish community in New York where a

Mexican domesticworker harbouring aTaenia solium tape-

worm had apparently contaminated the family’s food with

eggs from her tapeworm1.

T. solium is a cestode parasite that is transmitted between pigs and

people. Pigs may harbour the parasite’s larval stage in their muscles

(Figure 1). Ifwe accidentally eat oneof these larvae in poorly cooked

pig meat, the adult tapeworm will develop in our small intestine.

Humans are the only definitive host (the host in which the parasite

undergoes sexual reproduction) for this species of tapeworm. A

person with the tapeworm releases the parasite’s eggs in their

faeces, and the lifecycle is completed if foods contaminated with

these eggs, or indeed the faeces themselves, are consumed by a pig.

The disease is fully transmitted only where pigs roam freely and

humans defecate in areaswhere thepigs are free roaming. This does

nothappen inNewYork.However, it doeshappenacross large areas

of central and South America, Africa and east and south-east Asia.

In former times T. solium and the human brain disease that the

parasite causes, neurocysticercosis, were endemic throughout

Europe and other parts of the current-day First World; however,

improved public sanitation and hygienic standards for raising pigs

have seen thedisease in pigs eliminatedwithout any efforts directed

specifically to the disease. Likely the same will happen, in time,

through economic development in those areas of the world where

thedisease remains endemic today. Unfortunately, this is unlikely to

occur in our lifetimes. Meanwhile, those living in the T. solium

endemic areas of the world continue to suffer epilepsy and death

due to the presence of T. solium cysts in their brain. Keep in mind

that people from poor countries can travel anywhere with their

intestinal residents and deliver their tapeworm eggs to you or me,

just as happened in the New York community referred to above.

No point turning vego. Consider this: I have a worm and I am

not particularly hygienic when I go to the toilet. I make your salad;

bingo – see you in the neurology clinic.

T. solium is not an obscure parasite. The Food and Agriculture

Organization of the United Nations considers it to be the most

important foodborne parasitic infection from a global perspective2.

T. solium is the most frequent preventable cause of seizure dis-

orders, being associated with 29% of people with epilepsy3. The

World Health Organization list T. solium as one of 16 Neglected

Tropical Diseases, and is actively promoting efforts to reduce the

parasite’s transmission4.

For those living in poor countries in which T. solium is endemic,

help is at hand from, of all places, Australia. I say ‘of all places’

because the parasite is not, and has never been known to be,

endemic here. Nevertheless, a research program at the University

of Melbourne with an original genesis way back in the 1970s led

eventually to the development of both the first effective non-living

vaccine against a eukaryotic parasite and eventually also to a vaccine

that can stop pigs being infected with T. solium. The vaccine uses a

recombinant antigen known as TSOL18. Several independent ex-

perimental trials of TSOL18 have confirmed that it is extraordinarily

effective5. A field trial of the vaccine was undertaken in which pairs

of young piglets were distributed to families living in a T. solium

endemic region of north-east Cameroon6. One animal from each

pair was vaccinated and one acted as a control. When the animals

were of normal eating-age (~12 months), they were recovered

from the farmers and assessed for T. solium infection. About 20%

of the controls were infected but not a single parasite was found in

any of the 110 vaccinated animals.

So far, so good. However the field trial involved a single cohort of

animals that was vaccinated. In a real life situation, new disease

susceptible piglets are born into the community more-or-less

every day. Difficulties arise when we try to work out a feasible

and sustainable program to deliver vaccination regularly enough

to prevent pigs becoming infected on an on-going basis. The

‘standard’ vaccination protocol calls for two immunizations

about a month apart. It is difficult to give any veterinary care to

pigs in the communities where T. solium is transmitted; however

having to deliver two vaccines a month apart would be close to

impossible.

Under theMicroscope


Using knowledge about the parasite’s development in pigs and

information from the successful Cameroonian field trial, various

scenarios involving vaccination in pigs were compared for their

predicted effectiveness to control T. solium transmission7. One

scenario that appeared relatively practical was to vaccinate at

4-monthly intervals. While the scenario looked good theoretically,

it could not be recommended because we had no information

about whether the vaccine would raise a protective response if

the interval between primary and secondary injections was longer

than 4 weeks.

Recently we have completed an experiment in which pigs received

their secondary immunizationwithTSOL18at4,8,12,16or20weeks

after the first injection8. The results were very promising. Antibody

responses to the vaccine generally increased beyond the ‘standard’

4-week interval. Responses seen in the animals vaccinated at a

12 week interval were the best, and field evaluation of T. solium

interventions are about to begin in several endemic regions of Africa

that will involve vaccinations at 3 or 4 monthly intervals.

Despite the solid progress that has beenmade so far, much remains

to be achieved before we would be likely to see pig vaccination

contribute to reducing the incidence of neurocysticercosis in peo-

ple. One major difficulty with implementation of pig vaccination to

prevent transmission of T. solium is that the owners of the animals

often have little incentive to undertake control measures because

the infection does not often directly cause illness or death in pigs.

In the future, combination vaccines that include TSOL18 and also

antigens that can provide protection against pig deaths caused by

Classical Swine Fever (CSF) in the Americas, or African Swine Fever

(ASF) in African countries, would improve the acceptability of pig

vaccination by providing an economic incentive for the animal

owners to vaccinate. While the potential for a combination with

CSF is something that can be explored immediately because com-

mercial vaccines alreadyexist, there is yet tobeacommercial vaccine

for ASF.

As for the Jewish community in New York who use domestic staff

sourced from T. solium endemic countries, and of course all the

people who live permanently in those endemic countries, the risk

of exposure to T. solium remains. Hopefully we will be able to

overcome the practical difficulties around working with pigs in

T. solium endemic areas so that implementation of pig vaccination

can reduce T. solium transmission and decrease the incidence of

human neurocysticercosis as a result.

References1. Schantz, P.M. et al. (1992) Neurocysticercosis in an Orthodox Jewish community in

New York City.N. Engl. J. Med. 327, 692–695. doi:10.1056/NEJM199209033271004

2. Robertson, L.J. et al. (2013) Have foodborne parasites finally become a global

concern? Trends Parasitol. 29, 101–103. doi:10.1016/j.pt.2012.12.004

3. Ndimubanzi, P.C. et al. (2010) A systematic review of the frequency of neurocy-

ticercosis with a focus on people with epilepsy. PLoS Negl. Trop. Dis. 4, e870.


4. Organization, W.H. (2015) Investing to overcome the global impact of neglected

tropical diseases. Third WHO report on neglected tropical diseases. WHO/HTM/

NTD/2015.1.

5. Lightowlers, M.W. (2010) Eradication of Taenia solium cysticercosis: a role for

vaccination of pigs. Int. J. Parasitol. 40, 1183–1192. doi:10.1016/j.ijpara.2010.05.

001

6. Assana, E. et al. (2010) Elimination of Taenia solium transmission to pigs in a field

trial of theTSOL18 vaccine inCameroon. Int. J. Parasitol.40, 515–519. doi:10.1016/

j.ijpara.2010.01.006

7. Lightowlers, M.W. (2013) Control of Taenia solium taeniasis/cysticercosis: past

practices and new possibilities. Parasitology 140, 1566–1577. doi:10.1017/S003

1182013001005

8. Lightowlers, M.W. et al. (2016) Anamnestic responses in pigs to the Taenia solium

TSOL18 vaccine and implications for control strategies. Parasitology, in press.

BiographyMarshall Lightowlers has been a full-time research scientist

supported by medical research funding for more-or-less all of his

working life.He currently holds appointments as Laureate Professor

at The University of Melbourne’s Faculty of Veterinary and Agricul-

tural Sciences, and Principal Research Fellow with the NHMRC.

(a)

(b)

Figure 1. (a) Taenia solium cysts (cysticerci) in the tongue of a naturallyinfected pig (Photo: M. Donadeu). (b) Cysticerci in the muscles of aheavily infected pig.

Under theMicroscope


Seafood-borne parasitic diseases in Australia:how much do we know about them?

Shokoofeh Shamsi

School of Animal and VeterinarySciencesCharles Sturt UniversityWagga Wagga, NSW 2650, AustraliaTel: +61 2 6933 4887Email: [email protected]

Fish are host to many parasites, some of which can cause

disease inhumans.With the increase in cultural andculinary

diversity and the increased popularity of eating raw or

slightly cooked seafood dishes in Australia it is speculated

that seafood-borne parasitic infections in Australian consu-

mers may rise. Seafood-borne zoonotic parasites are recog-

nised as a significant public health concern worldwide.

In Australia there are few reports of infection in humans in

themedical literature. Australian Government enforcement

agencies rate the risk of seafood-borne zoonosis as low;

however, the prevalence of seafood-borne zoonoses may be

under-reported in Australia due to misdiagnosis. Although

food safety regulations and import controls for seafood

in Australia are strict, the focus is more on the control of

food-borne bacterial, viral and chemical contaminant rela-

ted illnesses rather than parasitic diseases.

Increasing demand for raw and exotic seafood has significantly

expanded the geographical and demographic limits of fish-borne

parasitic infections. Medical literature on the regular consumption

of seafood is plentiful. It is heavily promoted for prolonging life,

aiding in childhood development, increasing brain stimulation

and even to help our pets. However, there is an increasing risk of

contracting parasitic zoonoses from consuming raw and under-

cooked seafood, which should be of concern in Australia. Zoonotic

parasites are common in Australian seafood, and can pose a major

health risk to people who consume seafood or work in a fishing

industry1. Among zoonotic parasites in seafood, Anisakid nema-

todes are of the greatest significance due to their high prevalence in

wild caught fish2 and also due to the severity of the disease they

cause, which is known as anisakidosis. Other nematodes such as

Gnathostoma andAngiostrongylus aswell as platyhelminths suchas

Diphylobothrium, Clonorchis and Paragonimus, can occasionally

cause seafood-borne zoonotic infections3.

Infection in humans (zoonosis)Anisakidosis results from accidental infection with one or more

larvae of species of certain genera of anisakids, including Anisakis

spp, Contracaecum spp and Pseudoterranova spp. Infection with

larvae of Anisakis and Pseudoterranova is of common occurrence,

whereas infection with other genera, such as Contracaecum has

been reported less frequently. Humans usually become infected

with these parasites after eating raw, undercooked or improperly

processed fish or seafood (Figure 1). Clinically, several different

types of human anisakidosis have been described based on the

location of the parasite. This includes gastro-intestinal, visceral,

oropharyngeal and transient luminal anisakidosis4. The first two

types are associatedwith severe symptoms such as vomiting, severe

pain in lower abdomen and fever.

In recent years, it has been recognised that an allergic response can

occur in humans due to live anisakids or food in which worms have

been killed by cooking or pasteurisation5. Other allergic disorders,

such as chronic urticaria may also result from parasitism6. There is

anecdotal evidence in other countries (e.g. Spain) that some allergic

reactions to seafood are indeed allergic reaction to anisakids in

seafood.

The Australian scenarioIn Australia our knowledge about these important parasites is poor.

Although anisakid nematodes are commonly found in Australian

marine fish (Figure 2), such as mackerel, flathead, snapper and

whiting2,7,8, all forms of seafood are available and popular in Aus-

tralia and 40% of fish consumed raw in Australia have been consid-

ered as infected with Anisakis at the time of processing for trade1,

there are only a handful of documented confirmed cases acquired in

Australia.

Case 1: There has been only one documented confirmed case of

infection with anisakid nematodes acquired in Australia9. This case

involved a 41-year-old South Australian woman of Tongan descent

who became ill after eating raw mackerel that had been caught

locally. She was hospitalised with severe gastrointestinal pain,

diarrhoea and vomiting that progressively worsened for 3 weeks.

The diagnosis only occurred after a worm was passed in her faeces.

The initial presumptive identification of the larva was an intestinal

nematode, possibly a species of the Trichostrongylus or Ascaris

genera. However, on further detailed microscopic examination,

the larva was identified as a species of Contracaecum. Had the

worm not been found, or properly examined by a taxonomist,

Under theMicroscope


misdiagnosis would have occurred. It is very rare for anisakid larvae

to be found after passing through the gastro-intestinal tract as was

the case in this Australian patient.

Since then the author has come across several suspected cases of

anisakidosis, mostly among Australian travellers after returning

home (unpublished cases). The cases could not be confirmed due

to the lack of knowledge, clinical suspicion and a reliable diagnostic

technique in Australia. In many countries, the disease is usually

diagnosed by endoscopy, radiography, or surgery if the worm has

embedded within the gastro-intestinal tract in patients with symp-

toms and a history of consuming raw seafood. Serological tests to

detect parasite allergens have also been recently developed10.

However, inAustralia there is no standard test available for diagnosis

of these important parasites. General health practitioners are not

aware of the presence and high abundance of these parasites in

Australian fish and diagnostic laboratory staff are not trained to

Definitive hosts

1st intermediatehosts

2nd intermediate/paratenic hosts

Figure 1.General life cycle of anisakid nematodes. Adult nematodes inhabit stomachof definitive hosts, includingmarinemammals, piscivorousbirdsand large predatory fish in which they reproduce and lay eggs. Eggs pass through faeces to the water. Embryonated eggs or hatched larvae areingested by first intermediate hosts, including awide range of aquatic invertebrates. Larvae develop furtherwhen infected first intermediate hosts arepredated uponby second intermediate or paratenic hosts, including a broad range of fish species. The larval stages of Anisakids are not host specificand awide range of fish species can become infected as intermediate or paratenic hosts. This allows the parasite to bewidely distributed by passingthrough several fish species and infect awide rangeofmarinemammals and fish-eating birdswhere they complete their life cycle andbecomeadults.Thus infection with anisakids is not limited to one simple food chain but rather a wide network of species, increasing the impact and importance ofthese parasites. As shown in the picture humans become infected by consuming infected seafood (fish and invertebrates such as crustaceans).

Figure 2. Anisakid larvae (circled) on the surface of the internal organsof a fish caught in South Australia. If fish is not gutted immediatelyafter being caught these parasites migrate toward flesh of the fish(photograph by Shokoofeh Shamsi).

Under theMicroscope


identify these worms. As a result, the full extent of hidden cases of

the disease in Australia remains unknown.

Case 2: Another confirmed case report of fish-borne parasitic

disease in humans in Australia occurred in 201111. In this case, a

couple became infected withGnathostoma species after consump-

tion of ‘Black Bream’ (possibly Acanthopagrus berda or Hephaes-

tus jenkinsi), which they caught in the Calder River in Western

Australia. They presented with recurring skin swellings. The fish

had been cooked on a camp fire, but the cooking time was unclear.

The diagnosis was based on eosinophilia and a positive serology.

Humans become infected accidentally by consuming third-stage

larvae. Gnathostoma larvae are unable to mature inside the human

body, and instead migrate through visceral and cutaneous tissues,

causing a variety of generalised, non-specific symptoms. Gnathos-

tomiasis is generally endemic in parts of the world where seafood is

consumed raw, such as in South-East Asia and Japan, but also more

recently in LatinAmerica, India andAfrica, and in travellers returning

from these areas12.

Another aspect of Australia’s seafood consumption that must be

considered is the large volume of imported seafood that Australians

consume. The most important sources of seafood products are

Thailand, New Zealand, Vietnam and China (frdc.com.au/knowl-

edge/Factsheets/Factsheet_Imported_Seafood_in_Australia.pdf).

The literature describes several zoonotic parasites affecting fish

endemic to these countries13. CSIRO conducted a comprehensive

review of AQIS’s imported seafood testing protocols14 and con-

cluded that imported seafood does not pose any greater health risk

to the consumer than locally produced seafood. Nevertheless

proper precautions when preparing seafood in order to reduce

the risk of illness or disease has been recommended. However,

recommendations to eliminate or reduce parasites have not been

dealt with in details.

PreventionGastro-intestinal anisakidosis can beprevented byproperly cooking

fish to an internal temperature of approximately 638C or freezing at

or below�208C for 7 days15. Thiswillmost likely preventmost other

parasites as well.

Consuming raw seafood in reputable restaurants only, where chefs

are trained to recognise infected seafood.

ConclusionIn the absence of standard diagnostic techniques it is difficult to

have a realistic estimation of occurrence of seafood borne parasitic

diseases in the country. As Chai et al.12 stated it appears that fish

borne parasitic disease in Australia remains a ‘public health orphan’

with very little research having been carried out on what the real

risks are. The economic value of the fishing industry and health

of consumers must be protected. Therefore further research is

required to fill in the current knowledge gaps of the biology and

ecology of these parasites and the risk they pose to consumers and

workers.

References1. Anantanawat, S. et al. (2012) A semi-quantitative risk assessment of harmful

parasites inAustralianfinfish. SouthAustralian Research&Development Institute,

2012.

2. Shamsi, S. et al. (2011)Occurrence and abundance of anisakid nematode larvae in

five species of fish from southern Australian waters. Parasitol. Res. 108, 927–934.

doi:10.1007/s00436-010-2134-1

3. Butt, A.A. et al. (2004) Infections related to the ingestion of seafood. Part II:

parasitic infections and food safety. Lancet Infect. Dis. 4, 294–300. doi:10.1016/

S1473-3099(04)01005-9

4. Smith, J.W. (1999) Ascaridoid nematodes and pathology of the alimentary tract

and its associated organs in vertebrates, including man: a literature review.

Helminthological Abstracts 68, 49–96.

5. Audicana, M.T. and Kennedy, M.W. (2008) Anisakis simplex: from obscure

infectious worm to inducer of immune hypersensitivity. Clin. Microbiol. Rev.

21, 360–379. doi:10.1128/CMR.00012-07

6. Daschner, A. and Pascual, C.Y. (2005) Anisakis simplex: sensitization and clinical

allergy. Curr. Opin. Allergy Clin. Immunol. 5, 281–285. doi:10.1097/01.all.0000

168795.12701.fd

7. Doupe, R.G. et al. (2003) Larval anisakid infections of some tropical fish species

from north-west Australia. J. Helminthol. 77, 363–365.

8. Shamsi, S. et al. (2011) Mutation scanning-coupled sequencing of nuclear ribo-

somal DNA spacers (as a taxonomic tool) for the specific identification of different

Contracaecum (Nematoda: Anisakidae) larval types.Mol. Cell. Probes 25, 13–18.

9. Shamsi, S. andButcher, A.R. (2011) First report of human anisakidosis in Australia.

Med. J. Aust. 194, 199–200.

10. García, M. et al. (1997) The use of IgE immunoblotting as a diagnostic tool in

Anisakis simplex allergy. J. Allergy Clin. Immunol. 99, 497–501. doi:10.1016/

S0091-6749(97)70076-9

11. Jeremiah,C.J. et al. (2011)Gnathostomiasis in remotenorthernWesternAustralia:

the first confirmed cases acquired in Australia. Med. J. Aust. 195, 42–44.

12. Herman, J.S. and Chiodini, P.L. (2009) Gnathostomiasis, another emerging

imported disease. Clin. Microbiol. Rev. 22, 484–492. doi:10.1128/CMR.00003-09

13. Chai, J.-Y. et al. (2005) Fish-borne parasitic zoonoses: status and issues. Int.

J. Parasitol. 35, 1233–1254. doi:10.1016/j.ijpara.2005.07.013

14. Moir, C. (2009) Review of the current testing protocols for imported seafood

products. CSIRO Report reference number R-661-03-11. 90 pp.

15. Hochberg, N.S. and Hamer, D.H. (2010) Anisakidosis: perils of the deep. Clin.

Infect. Dis. 51, 806–812.

BiographyDr Shokoofeh Shamsi is a senior lecturer in veterinary Parasito-

logy. After completing a Masters Degree in Medical Parasitology

(Tehran University of Medical Sciences) she gained her PhD in

Veterinary Parasitology from The University of Melbourne. She is a

taxonomist who couples conventional morphological techniques

with modern molecular technologies to diagnose infections and

identify parasite species, especially aquatic parasites. This has

resulted in the discovery of several new pathogenic species and

strains, and their recognition as disease causing agents in humans

and animals in Australia and worldwide.

Under theMicroscope


Children, snails and worms: the Brachylaimacribbi story

Andrew R Butcher

SA PathologyMicrobiology and InfectiousDiseasesSA, AustraliaEmail:[email protected]

Brachylaimids are parasitic trematode fluke worms that

have a terrestrial life cycle involving land snails and slugs

as thefirst and/or second intermediate hosts for the cercarial

and metacercarial larval stages. A wide range of mammals,

birds, reptiles andamphibians are thedefinitivehosts for the

adult worm. Brachylaima spp. have been reported from

most continents including Europe, Africa, Asia, North and

South America and Australia. There are over 70 described

species in the genus with seven species indigenous to Aus-

tralia. Although Brachylaima spp. are a cosmopolitan ter-

restrial trematode they have not been recorded to infect

humans other than the three Brachylaima cribbi infections

reported in two children and an adult from South Australia.

The publication of a new species of brachylaimid, Brachylaima

cribbibyButcher andGrove in2001was the culminationof a 12-year

scientific journey that would not have been possible without a

broad scientific network of colleagues and some ‘scientific luck’,

which enabled the establishment of a laboratory life cycle using

parasite eggs recovered from the stool of an infected human1.

My long-term association, as an active member of the Australian

Society for Microbiology and the special interest groups played a

significant role in having a network of colleagues to help find

techniques and data to assist in the study of this parasite.

The publication of the first human Brachylaima sp. infections

followed the detection of a fluke worm egg, in the stool of two

South Australian children, who had never travelled overseas. The

eggs were identified as belonging to a genus of trematode worm

endemic in the local area2. After conversations with many local and

interstate colleagues, a veterinary parasitology colleague Michael

O’Callaghan remembered the post-doctoral work of Thomas Cribb

in South Australia. He had studied and published work on a

brachylaimid in South Australia that infected mice3,4. This provided

the data on a local trematode that had an eggmorphologymatching

the eggs detected in the children’s stools.

However, was this a true human infection that resulted in the

establishment ofmature gravidworms producing the eggs detected

in their stool or was it a spurious infection? The answer to this

questionwas resolved 18months laterwhen anelderly lady from the

mid-north of South Australia presented with chronic diarrhoea.

Microscopic examination of her stool detected Brachylaima eggs,

which matched the morphology of the eggs detected in the two

children. In addition, on the first day following treatment with

praziquantel a gravid degenerate adult Brachylaima worm was

recovered from her stool5. This provided the evidence that a

Brachylaima sp.was infecting humans and completing its life cycle.

(a) (b)

Figure 1. (a) European helicid and hygromiid land snails aestivate over summer to escape the ground heat by attaching to fence posts,which presentsan easy meal for natural definitive hosts like birds. (b) Fertile Brachylaima cribbi egg being smooth shelled with an inconspicuous operculum,an abopercular knob or thickening and measuring 26–32mm (29.1mm) long and 16–17.5mm (16.6mm) wide.

Under theMicroscope


However, due to the advanced state of decomposition of the worm

there was insufficient morphological detail to establish the species

identification.

Previous to the finding of the third case described above I had

commenced the cultivation of introduced European helicid and

hygromiid land snails (Figure 1a) in homemade terrariums using

techniques published for snail farming6. My office in the diagnostic

Microbiology laboratory had now become part snail culture labo-

ratory. These snailswereused to study the larval stages in an attempt

to understand how humans could have become infected. At the

same time I had obtained a copy of an article by Mas-Coma and

Montoliu detailing the life cycle of Brachylaima ruminae from the

duodenumofa rodent fromtheMediterranean islandofFormentera

(Spain)7. This article provided methods for the establishment of a

laboratory life cycle. With these methods in hand and my snail

cultures thriving I set myself the task to attempt to establish a

laboratory life cycle using eggs recovered from the stool of the third

patient. Faecal sediment of the patient’s stool containing washed

concentrated eggs was smeared on wet filter paper and was fed to

laboratory snails in Petri dishes overnight. Eight weeks later, when

the snails were exposed to moisture in a Petri dish, cercariae by

the thousands were observed emerging from the snails (Figure 2).

Cercariae were collected in water and used to infect the common

brown garden snails Cornu aspersum (Helix aspersa) via the

respiratory orifice. After 8–10 weeks mature metacercariae were

harvested from kidneys of the snails. Mice were inoculated orally

withmetacercariae. Sevenweeks later eggswere detected inmouse

(a)

(b) (c)

Figure 3.Brachylaimacribbi adult worm. (a) Oral (os) and ventral suckers (vs) and genital pore (gp). Bar = 500mm. (b) Ventral viewof the oral and ventralsuckers showing rows of tegumental spines. Bar = 100mm. (c) Genital pore showing extended cirrus (c) releasing sperm (s). Bar = 100mm.

Figure 2. Scanning electron microscopy ventral view of a Brachylaimacribbi cercaria showing oral (os), ventral suckers (vs) and elongatedsensory papilla (sp). Bar = 50mm.

Under theMicroscope


faeces. Gravid adult Brachylaima worms were recovered from the

small intestine for characterisation andmorphological identification

(Figure 3). Eggs frommouse faeces and dissectedwormswere used

to continue the life cycle (Figure 4) in the laboratory to study the life

cycle kinetics.

At the same time as I was using my spare time to tinker with this life

cycle experiment the former Institute of Medical and Veterinary

Science (IMVS) was undergoing amalgamation with the Queen

Elizabeth Hospital (TQEH) Laboratories. Professor David Grove

was the Clinical Microbiologist Head of TQEH Microbiology

Department and a world recognised expert clinical parasitologist.

At that time Professor Grove offeredme the opportunity to join the

TQEH Microbiology team under the amalgamation program to

continue the Brachylaima study and commence a PhD under his

supervision. That is scientific luck, right place at the right time and

the rest is now history, with the following years being the most

simulating and productive science of my career.

(e)

(d)

(a)

(b)

(c)

Figure 4. Life cycle of Brachylaima cribbi. The definitive host sheds eggs (a) in their faeces. The egg is eaten by introduced European land snails(b), which hatch in the snail’s gut to release themiracidium. Themiracidiumdevelops into a sporocyst in the snail’s digestive gland.Mature cercariae(c) released from the sporocyst emerge from snail passing into the environment. Cercariae infect other land snails and develop into metacercariae(d) in the snail’s kidney. The definitive host (mammals, birds and reptiles) (e) ingest infected snails, releasing metacercariae, which migrate to thesmall intestine, attach and develop into mature adult worms. Humans are incidental definitive hosts being infected after eating raw snails containingmetacercariae.

Under theMicroscope


Human cases

Since the publication of the three human cases to my knowledge

there have been a further 12 laboratory confirmedhuman infections

(A.R. Butcher and D.I. Grove, unpublished observations). Part of

my PhD studies was the follow-up of all human infections with a

standard questionnaire to obtain clinical history. The main present-

ing symptoms were mucoid, watery diarrhoea, abdominal pain,

anorexia and weight loss or poor weight gain in children. All

identified cases were in patients who lived in rural or semi-rural

districts where helicid and hygromiid land snails infected with

B. cribbimetacercariae were abundant. The likely infection source

was the ingestionof rawsnails. Themajorityof infections (80%)were

in children under the age of two years. At this age the mouthing of

objects is a common behavioural characteristic with parents of the

infected children detailing their child’s consumption of snails. For

the three adults infected they had all eaten home-grown garden

vegetables contaminated with snails. The duration of symptoms

ranged from 1 month to 2 years with the majority of infections

diagnosed within 1–2 months of the first signs of symptoms. All

patientswere treated successfully with praziquantel with no adverse

side-effects. The most commonly used dose in both adults and

children was 20 mg/kg once per day for 3 days.

Laboratory diagnosis

Diagnosis of the infection relies on the detection of typical eggs in

the faeces which are asymmetrical, being ovoid, with one side

slightly flattened. They have a smooth shell, inconspicuous oper-

culum, an abopercular knob or thickening and measure 26–32mm

by 16–17.5mm. The eggs generally contain a well-developed mira-

cidium (Fig. 1b). However, in chronic infections many of the eggs

may be infertile, being of similar morphology but smaller size and

lacking a developed miracidium. Infected land snails have been

reported from coastal and inland Western Australia, South Australia

and western Victoria8. Therefore, the potential exists for further

human infections, especially in children living in these regional

districts.

References1. Butcher, A.R. and Grove, D.I. (2001) Description of the life-cycle stages of Brachy-

laima cribbi n. sp. (Digenea: Brachylaimidae) derived from eggs recovered

from human faeces in Australia. Syst. Parasitol. 49, 211–221. doi:10.1023/A:1010

616920412

2. Butcher, A.R. et al. (1996) Locally acquired Brachylaima sp. (Digenea: Brachylai-

midae) intestinal fluke infection in two South Australian infants.Med. J. Aust. 164,

475–478.

3. Cribb, T.H. (1990) Introduction of a Brachylaima species (Digenea: Brachylaimi-

dae) to Australia. Int. J. Parasitol.20, 789–796. doi:10.1016/0020-7519(90)90013-D

4. Cribb, T.H. and O’Callaghan, M. (1992) An unusual trematode infecting domestic

chickens. Aust. Vet. J. 69, 69–70. doi:10.1111/j.1751-0813.1992.tb07456.x

5. Butcher, A.R. et al. (1998) First report of the isolation of an adult wormof the genus

Brachylaima (Digenea: Brachylaimidae), from the gastrointestinal tract of a

human. Int. J. Parasitol. 28, 607–610. doi:10.1016/S0020-7519(97)84372-X

6. Baker, G.H. (1991) Production of eggs and young snails by adult Theba pisana

(Müller) and Cernuella virgata (da Costa) (Mollusca: Helicidae) in laboratory

culture and field populations. Aust. J. Zool. 39, 673–679. doi:10.1071/ZO9910673

7. Mas-Coma, S. andMontoliu, I. (1986) The life cycle of Brachylaima ruminae n. sp.

(Trematoda: Brachylaimidae), a parasite of rodents. Z. Parasitenkd. 72, 739–753.

doi:10.1007/BF00925095

8. Butcher, A.R. and Grove, D.I. (2003) Field prevalence and laboratory susceptibility

of southern Australian land snails to Brachylaima cribbi sporocyst infection.

Parasite 10, 119–125. doi:10.1051/parasite/2003102119

Biography

Dr Andrew Butcher is a retired medical scientist with 38 years

experience as a diagnostic medical microbiologist with a special

interest in parasitology. He has been an active member of the ASM

Parasitology and Tropical Medicine Special Interest Group for

many years being national convenor for 7 years. He has been

involved in teaching diagnostic medical parasitology and one

of the founding committee members of the ASM Parasitology

Master Class.

Under theMicroscope


Malaria: global challenges for malaria eradication

Graham Brown

The University of MelbourneVic. 3010, AustraliaEmail: [email protected]

Stephen Rogerson

The University of MelbourneVic. 3010, AustraliaEmail: [email protected]

Theenormousdecline in theannualmorbidity andmortality

frommalaria is the spectacular global public health success

of the past decade. This achievement results largely from

increased finance for investment in measures known to

prevent malaria: bednets treated with long-lasting insecti-

cides, chemoprophylaxis, and rapid access to effective treat-

ment.Suchhasbeenthesuccessof thesemeasures thatplans

are being put in place to achieve the vision of a malaria-free

world within the next three decades. Large financial and

political commitments and ongoing research will be re-

quired to maintain the gains, overcome known and un-

known challenges such as drug and insecticide resistance,

and to achieve those goals. Effective vaccines ormethods for

reducingmosquito vectorial capacity would add enormous-

ly to the chance of achieving this goal. The aim of this article

is to summarise the current status of malaria control, the

recent research successes, the challenges being addressed,

and the plan for progress to elimination of malaria in the

longer term.

Spectacular progress in reducing the burden

of malaria

Funding for major efforts in implementation of known effective

preventive measures in the past 15 years has caused an almost 40%

reduction in incidenceofmalaria disease episodes. Thesemeasures,

together with increased access to effective treatment, have led to

a reduction in malaria death rates of 60% globally, and 71% in

children under five1. It is estimated that between 2001 and 2015,

~6.2million lives were saved by increasing the provision of these

services. Approximately 110 countries are free from malaria, ~40

have committed to an elimination timetable, while 70 focus

predominantly on malaria control. Many factors contributed to this

success from the many constituencies that make up the Roll Back

Malaria Partnership2, but most of the progress in Africa is usually

attributed to widespread distribution of bednets treated with long-

lasting insecticides, combined with increased access to highly-

effective drugs3.

The current situation

According to the latest information provided by the WHO, ~1.2

billion people are at risk ofmalaria with themajor burden of disease

carried by young children and pregnant women living in endemic

areas.1

The burden of malaria is not evenly distributed across the world.

About 90% of malaria deaths occur in sub-Saharan Africa and recent

estimates suggest that 80% of malaria deaths occur in only 15

countries. The largemorbidity resulting fromP. vivax is seenmainly

in the Asia Pacific region. Further general information on malaria is

available in themost recentWorldMalariaReport fromWHO1.There

has been great recent interest in the finding that infections with the

simian malaria P. knowlesi have become increasingly important in

Malaysia and surrounding countries.

The strategy for malaria control leading

to elimination

The overall guiding strategy for addressingmalaria is summarised in

theGlobal Technical Strategy forMalaria Elimination 2016–2030,

which was approved and adopted by the World Health Assembly in

May 20154. The strategy calls for accelerated action towards malaria

elimination in countries and regions, but does not set a time frame

for global eradication. Notably this document recognises the con-

tinuum of activities from malaria control to elimination, the

Under theMicroscope


importance of amulti-disciplinary approach, and the need for tailor-

made strategies for different situations, even within one country, in

other words, ‘One size does not fit all’. One area, for example a

mountainous region, may have only the problem of introduced

malaria occurring in people of all ages, whereas another highly

endemic area may need to focus on children and pregnant women

who remain at risk and suffer the consequences all year round.

The Global Malaria Programme of WHO provides support to coun-

tries, compiles the Annual Global Malaria Report, and through the

Malaria Policy Advisory Committee provides a normative role in

reviewing new data or new problems to provide advice for policy

implementation at country level.

This WHO strategy is complemented by the Roll Back Malaria

advocacy plan, Action and Investment to Defeat Malaria

2016–2030, that was approved in 20155. The publication outlines

both the health and economic benefits that wouldflow frommalaria

elimination, and like the Global Technical Strategy was the result of

extensive consultative processes involving the participation ofmore

than 400 malaria experts from 70 countries. Ambitious but achiev-

able global targets were established, including:

* reducing malaria case incidence by at least 90% by 2030* reducing malaria mortality rates by at least 90% by 2030* eliminating malaria in at least 35 countries by 2030* preventing a resurgenceofmalaria in all countries that aremalaria-free.

The timeline of 2016–2030 is aligned with the 2030 Agenda for

Sustainable Development, the new global development framework

adopted by all UN Member States in September.

Malaria elimination

The major progress already referred to is the result of implemen-

tation of known effective preventive measures, including vector

control with long-lasting insecticide-treated nets and indoor resid-

ual spraying; chemoprophylaxis including intermittent preventive

treatment of malaria in pregnancy (IPTp); intermittent preventive

treatment of malaria in infants (IPTi) or malaria chemoprevention

during the season of malaria transmission (SMC); and provision of

ready access to case management with rapidly-effective drugs,

primarily artemisinin combination treatments (ACTs).

As countries, or areas within countries, move towards elimination,

the emphasis changes from population level data and disease

control to intense surveillance for identification of individual infec-

tions, followed by planning and, most importantly, implementation

of a response. For example, teams might test everyone in a com-

pound or a village surrounding a person presenting with illness,

apply control measures locally, and search for the source of intro-

duced cases.

MalERA (Malaria Eradication Research Agenda)

Following the call for malaria eradication, the Bill & Melinda Gates

Foundation recognised the need for new and better tools for

elimination, andnewandbetter systems toguidedeliveryof services

to achieve those goals. Following extensive discussions by working

groups, a research agenda was developed (MalERA)6 that included:

* Better and safer drugs for ease of administration to populations.The ‘dream’ product profile is ‘SERCAP’, (Single EncounterRadical Cure And Prophylaxis), i.e. a single dose treatment to killnot only asexual blood-stage parasites but also liver and sexualstages. An intermediate goal would be an alternative and moreeffective drug to kill hypnozoites, the dormant liver stage respon-sible for relapse in P. vivax, but without the troublesome side-effects of primaquine, the only drug currently available for thatpurpose.

* Basic researchenablingcultureof all life cycle stagesof all parasitesinfecting humans would be required for development of vaccinesand for production of better drugs for eradication of hypnozoites.This would entail investment in culture of hepatic stages ofP. vivax, and achieving the dream of laboratory culture of spor-ozoites from gametocytes.

* The emphasis on elimination draws attention to P. vivax, com-mon outside Africa, and with separate challenges such as long-term relapses from hypnozoites, and transmission by vectorsbiting outdoors and during the daytime, which are therefore notamenable to the standard interventions for vector control.

Many considered that elimination would not be achieved without a

major breakthrough such as effective vaccines to reduce morbidity

and transmission, or a technology for vast reduction in vectorial

capacity of Anopheles gambiae, the most efficient and important

vector of P. falciparum in Africa.

It is a source of optimism that since the MalERA agenda was

proposed, fundingwasdirected to address important gaps, aMalaria

Eradication Scientific Alliance (MESA) was established, and now

through its MESA Track database tool can capture relevant research

anddevelopment projects. By the endofOctober 2015, 685projects

from 647 institutions and 84 countries had been documented7.

MESA is currently coordinating several expert groups to perform a

5-year ‘refresh’ of the MalERA agenda.

Progress is being made onmany other fronts, including conference

reports of the in vitro culture of sporozoites from gametocytes;

successful culture of hepatic stages of plasmodia; and development

of new antimalarial drugs, through ‘Medicines for Malaria Venture’

(MMV), a highly successful not-for-profit public-private partnership.

The timeframe for initial testing of newdrugs (or vaccines) has been

dramatically shortened by the development of a blood stage chal-

lenge model by Queensland-based researchers.

Under theMicroscope


The challenges for malaria control

and elimination

See Table 1 for a more comprehensive list. Some of the biological,

societal and economic requirements for elimination are listed in

Table 2.

Artemisinin resistance

The time andmodeof action of artemisinin are notwell understood,

and in vitro tests are not sufficiently developed and standardised to

replace the requirement for in vivo testing for the presence of

resistance.Awarning sign, beforeemergenceof clinical resistance, is

the detection of delayed parasite clearance from the blood over

the first few days of treatment. Because of the fear of emergence

of resistance, combination therapy is essential, for example with

piperaquine or lumefantrine, so the efficacy of partner drugs is also

of great importance to ensure complete parasite clearance. Recent-

ly, drug resistance has been associated with some but not all

mutations of the kelch gene in P. falciparum that will probably

assist in mapping resistance8,9. Melbourne-based researchers have

developed synthetic artemisinin-like compounds, which are prob-

ably not susceptible to kelch-based resistance.

Insecticide resistance

Chemical resistance to insecticides is a well known threat. Strong

movements against DDT that is cheap and effective for indoor

residual spraying (because of past evidence of harmful environmen-

tal effects when very widely distributed in agriculture) mean that

there are few suppliers globally, and alternatives are very expensive.

Behavioural resistance is another well known complication of the

introduction of effective vector control directed at mosquitoes that

bite indoors in the evening. Repeated studies have documented the

gradual increase in proportion of mosquitoes biting earlier in the

evening or later in themorning and resting outdoors where they are

not susceptible.

Another huge challenge, particularly for control of P. vivax is that

the majority of vectors for this parasite bite outdoors and during

the daytime. No satisfactory method for long-term control of these

vectors is available.

Elimination as an approach to drug resistance

in the Mekong region

Where artemisinin has been widely used in the Mekong region,

delayed parasite clearance has been noted inmany countries, and it

is felt that in the absence of alternative drugs, the only appropriate

approach is to eliminate malaria in the area where resistance has

already been detected. Pilot studies have been initiated to test the

feasibility of this approach10.

Fortunately, delayed parasite clearance due to artemisinin resis-

tance has not yet been seen in parasites from India and Africa, but

in the recent past, development of resistance to chloroquine

or sulphadoxine/pyrimethamine accompanied by ongoing use of

those drugs led to many unnecessary deaths.

Table 1. Some of the challenges for malaria control and elimination.

Parasite and vector factors

Artemisinin resistance (and resistance to partner drugs)

P. vivax hypnozoites

Detecting infections at low parasitaemia

Vector resistance (chemical and behavioural)

Substandard and counterfeit drugs

Societal and economic

Inadequate health services to deliver interventions

Lack of financial commitment

Lack of political will

Cross border coordination

Underserved minorities at high risk

Civil unrest

Table 2. Preparing for elimination.

Biological approaches

SERCAP a drug for ‘Single Encounter Radical Cure And Prophylaxis’

Highly sensitive non-invasive point-of-care tests

Highly effective malaria vaccines

Novel vector control strategies

Societal and economic

Political will for a long-term program at high cost per case

Financial commitment

Experienced work force

Technical and operational capacity

Ongoing research

Under theMicroscope


Funding shortfall

The major advances of the past two decades are at great risk from

lack of funding. Not only could progress stall, but major epidemics

of malaria could return to cleared areas, as has been seen so often

in the past11, with serious effects on people of all ages whose

immunity has declined through lack of boosting by recurrent

sub-clinical infections.

The case is now strong for the economic benefits that will follow

from the elimination of malaria5. This message needs to reach the

highest levels of government to ensure financial and political

commitment and the engagement not just of the health system,

but all sectors of society.

Strengthened Health Services

As themalariamapshrinks, the lastpocketsof transmission areoften

found in marginalised poor or rural populations with least access to

services. As episodes become infrequent, success will depend on a

general health service with the capacity to recognise and diagnose

infections, then respond with appropriate increased surveillance

to detect the origin of the infection and interventions to prevent

further spread.

Regional initiatives

APMEN Recognising the need for a partnership across regions and

across borders, the Asia Pacific Malaria Elimination Network was

establishedwith the supportof theAustraliangovernment in2008 to

facilitate collaboration amongst nations of the region that hadmade

commitments to achieve elimination within a defined time period.

This network enables regional collaboration and facilitates capacity

building and knowledge transfer among nations of the region that

share similar challenges such as artemisinin resistance, inadequate

finance for malaria, or a vast market penetration of counterfeit

drugs.

AICEM, the Australian Initiative for Control and Elimination of

Malaria, is funded by the Australian government to strengthen

malaria control in the Solomon Islands and Vanuatu, and at the

same time establish pilot projects for elimination of malaria in

certain island populations.

APLMA (www.aplma.org), the Asia Pacific Leaders Malaria Alli-

ance, co-chaired by the Heads of Government of Australia and

Vietnam, modelled loosely on the African Leaders Malaria Alliance,

aims to bring together Heads of State, politicians, and opinion

leaders of civil society to advocate for political and financial com-

mitment to the goals of elimination of malaria. Special task forces

are designed to address critical issues that could be barriers to

achievement of elimination such as financing, or the safety and

quality of medicines (to tackle the problem of counterfeit drugs

and ensure that only certified effective products are available in the

public andprivate sectors). Recently, theAllianceendorsedaMalaria

Elimination Roadmap towards a malaria-free Asia Pacific by 2030.

Elimination to eradication

The malaria eradication campaign of 50 years ago succeeded in

eliminatingmalaria frommanycountriesbutdidnot achieve its goals

in areas of most intense transmission such as heartland Africa.

However, many important lessons were learned about the need

for community engagement, political will, intensive surveillance as a

response, and the need for continued vigilance to prevent return of

malaria to a now susceptible entire population that has benefited

from a few years of interrupted transmission. As transmission

declines and malaria-specific services are phased out, it is critical

for awareness of the disease in all itsmanifestations to be retained at

a high level in general services for detection, management, surveil-

lance and response to any new episodes.

Lessons of the eradication campaign are being refreshed, for ex-

ample on mass drug administration to entire populations (MDA),

and comparing this strategy with treatment only of those found

with highly sensitive tests to be infected, thus avoiding the unnec-

essary use of drugs that could have side-effects, so called Mass

Screening and Treatment, or MSAT. Research on application of

moremodern tools such as rapid diagnostic tests, including nucleic

acid amplification or data transmission by mobile phones are

providing further data to guide strategy. For example, it is important

to decide whether the cost of a more sensitive but more expensive

and technically difficult nucleic acid detection test adds significant

benefit in elimination campaigns, or how PCR-based parasite

‘barcoding’ (genetic epidemiology) can be used to provide rapid

analysis of the source of emergent infections, the spread of drug

resistance, or the risk of the parasite’s appearance in susceptible

areas.

Othermajor lessons of the eradication campaignwere that research

must continue throughout the programs to deal with new chal-

lenges, and that every countryneeds todevelop thehuman resource

capacity to deal with the multi-disciplinary approach to this grand

challenge.

Recent vaccine breakthroughs; progress

with P. falciparum but little progress with

other species

Successful Phase 3 field trials of RTS,S vaccine in African

children. Development of the RTS,S vaccine, which provides

Under theMicroscope


35% efficacy against clinical episodes of P. falciparummalaria, was

an important milestone for the malaria vaccine field. The vaccine

includes a fusion protein containing the immunogenic repeat

regions of the circumsporozoite protein, T-cell epitopes of

the same molecule and the hepatitis B surface antigen that is

co-expressed with free hepatitis B virus surface antigen leading to

formation of a viral-like particle known as RTS,S. In recent very large

trials, participants had good access to malaria prevention and rapid

access to treatment, somortalitywas lowandnoeffect of vaccination

on mortality could be detected. There was a concerning, but

unexplained, slight increase in episodes of meningitis in the RTS,S

arm, requiring further evaluation. A recent review of RTS,S byWHO

recommended that the next step should be to initiate 3–5 pilot

implementation studies12. Further analysis of trial data showed that

the vaccine had higher efficacy against strains homologous to the

vaccine, suggesting both a strain-specific as well as a strain-inde-

pendentmechanism13. This vaccine is a very importantfirst step but

on its own, is unlikely to be the solution that is required.

Attenuated sporozoite vaccines

The most impressive of all malaria vaccine trial results are those

achieved with intravenous inoculation of radiation-attenuated

whole sporozoites in which up to 100% protection has been

achieved in Phase 1 studies using multiple doses of vaccine14. Field

studies in Mali and Tanzania have also demonstrated efficacy and

further trials are planned to optimise dose and delivery schedule14.

Conclusion

With the spectacular reduction of the burden of malaria in the past

15 years, the time is ripe to re-double efforts both to prevent

resurgence and to increase resources with the goal of achieving

the health and economic benefits that would result from disease

elimination. Major progress is being reported in developing the

tools to add to current successful interventions from basic research

into biology of hypnozoites and transition of gametocytes to spor-

ozoites, to field trials of vaccines and innovations in surveillance and

epidemiology. A multi-disciplinary response coupled with strong

community engagement, and ongoing political and financial sup-

port will be required to maintain the current rate of decline in

malaria, and hopefully achieve the ambitious goal of elimination.

References1. World Health Organization (2015) Malaria: fact sheet no. 94. http://www.who.int/

mediacentre/factsheets/fs094/en/

2. World Health Organization (2016) Roll back malaria partnership. http://www.

rollbackmalaria.org

3. Bhatt, S. et al. (2015) The effect of malaria control on Plasmodium falciparum in

Africa between 2000 and 2015. Nature 526, 207–211. doi:10.1038/nature15535

4. World Health Organization (2015) Global technical strategy for malaria 2016–

2030. http://www.who.int/malaria/publications/atoz/9789241564991/en/.

5. World Health Organization Secretariat, on behalf of the Roll Back Malaria Pro-

gramme. (2015)Action and investment todefeatmalaria 2016–2030. For amalaria-

free world. http://www.rollbackmalaria.org/about/about-rbm/aim-2016-2030

6. (2011) malERA – a research agenda for malaria eradication (PLoS medicine

monographic volume). http://collections.plos.org/malera?issue=info:doi/10.1371/

issue.pcol.v07.i13

7. Malaria Eradication Scientific Alliance (2015) MESA track. http://www.malariaer-

adication.org/mesa-track

8. Ariey, F. et al. (2014) A molecular marker of artemisinin-resistant Plasmodium

falciparum malaria. Nature 505, 50–55. doi:10.1038/nature12876

9. Ashley, E.A. et al. (2014) Spread of artemisinin resistance in Plasmodium

falciparummalaria. N. Engl. J. Med. 371, 411–423. doi:10.1056/NEJMoa1314981

10. World Health Organization (2015) Draft strategy for malaria elimination in the

GreaterMekong subregion (2015–2030). http://www.who.int/malaria/areas/great-

er_mekong/consultation-elimination-strategy/en/

11. Cohen, J.M. et al. (2012) Malaria resurgence: a systematic review and assessment

of its causes. Malar. J. 11, 122. doi:10.1186/1475-2875-11-122

12. World Health Organization (2015) Pilot implementation of first malaria vaccine

recommended by WHO advisory groups. http://www.who.int/mediacentre/news/

releases/2015/sage/en/

13. Neafsey, D.E. et al. (2015) Genetic diversity and protective efficacy of the RTS,S/

AS01 malaria vaccine. N. Engl. J. Med. 373, 2025–2037. doi:10.1056/NEJMoa

1505819

14. Richie, T.L. et al. (2015) Progress with Plasmodium falciparum sporozoite

(PfSPZ)-based malaria vaccines. Vaccine 33, 7452–7461. doi:10.1016/j.vaccine.

2015.09.096

Biographies

Graham Brown, AM Professor Emeritus at The University of

Melbourne, currently serves as the Deputy Chair of the Board,

and Chair of the Executive Committee of Roll Back Malaria. As a

clinician-researcher, his interests include malaria vaccine develop-

ment, antigenic variation and clinical infectious diseases.

Stephen Rogerson is a Professor at The University of Melbourne.

His research interests include the pathogenesis of malaria in preg-

nant women and young children, and tools for prevention of

malaria in these high-risk groups.

Member of Australian Society for Microbiology (MASM)

Want to upgrade from an associate to a full member?No points or cash required and only one supporting referee needed.Go to the ASM website, download and complete the form, and forward it with a CV to the NationalOffice.We will do the rest.

Under theMicroscope


Plasmodium knowlesi: an update

Balbir Singh

Malaria Research CentreUniversity Malaysia Sarawak94300 Kota SamarahanSarawak, MalaysiaTel: +6 082 581000Fax: +6 082 665152Email: [email protected]

There were only four species of Plasmodium that were

thought to cause malaria in humans until a large number

of human infections by Plasmodium knowlesi, a malaria

parasite typically found in long-tailed and pig-tailed maca-

ques,werereported in2004 inMalaysianBorneo.Since then,

cases of knowlesi malaria have been reported throughout

South-east Asia and also in travellers returning from the

region. This article describes the molecular, entomological

and epidemiological data which indicate that P. knowlesi is

an ancient parasite that is primarily zoonotic, and there are

three highly divergent sub-populations. It also describes the

detectionmethods forP. knowlesi, which ismorphologicaly

similar to P. malariae, and the clinical features and treat-

ment of this malaria parasite that is potentially fatal.

Malaria parasites and discovery of large focus

of human knowlesi malaria cases

Malaria is caused by parasites that belong to the genus Plasmodium

and there are more than 150 species of Plasmodium that infect

reptiles, birds andmammals1. These parasites, in general, tend to be

host-specific. Long-tailed and pig-tailed macaques (Macaca fasci-

cularis and M. nemestrina respectively) are hosts to five species

(P. knowlesi, P. inui, P. cynomolgi, P. fieldi and P. coatneyi). Only

four species of Plasmodium, namely P. falciparum, P. vivax,

P. malariae and P. ovale, were thought to causemalaria in humans

until a large number of human cases due to P. knowlesi were

reported in Sarawak, Malaysian Borneo over 11 years ago2. The

study in Kapit was prompted by observations that cases diagnosed

by microscopy as P. malariae had high parasitaemias, required

hospitalization and that 95% of patients were adults. This was in

contrast to P.malariae infections which typically are asymptomatic

with low parasitaemia and occur in all age groups. When blood

samples from 208 malaria patients at Kapit Hospital were analysed

by PCR assays, none were identified as P. malariae, although 141

had been diagnosed as P. malariae by microscopy. Fifty-eight

percent (120) were either single P. knowlesi infections or mixed

infections of P. knowlesi with P. falciparum and P. vivax. Misdi-

agnosis had occurred because the blood stages of P. knowlesi and

P. malariae are morphologically indistinguishable3.

Epidemiology and risk factors of acquiring

knowlesi malaria

Human infections with P. knowlesi have been reported throughout

Malaysia and in Thailand, Singapore, the Philippines, Vietnam,

Cambodia, Indonesia, Brunei, Myanmar and in the Nicobar and

Andaman Islands of India4,5. InMalaysia, P. falciparum and P. vivax

cases have declined over the pastfive years and P. knowlesihas now

become the most common cause of human malaria6,7. The true

incidence of knowlesi malaria is not known in other parts of South-

east Asia since not many large-scale studies have been undertaken

with molecular detection assays.

The geographical distribution of human P. knowlesi infections is

similar to that of the natural hosts ofP. knowlesi, the long-tailed and

pig-tailed macaques8. Reports from 1931 to 1970 identified maca-

ques as hosts of P. knowlesi in Peninsular Malaysia, Singapore and

the Philippines9, and a banded leaf monkey (Presbytis melalophos)

in Peninsular Malaysia9. Since 2007, P. knowlesi infections detected

by molecular methods have been described in macaques in Penin-

sular Malaysia, Malaysian Borneo, Singapore and Thailand4.

The transmission of P. knowlesi in nature has been shown to be

restricted to mosquitoes belonging to the Anopheles leucosphyrus

group10. The members of this forest-dwelling group of mosquitoes

that have been identified as vectors include An. latens (in Sarawak,

Malaysian Borneo)11, An. balabacensis balabacensis (in Sabah,

Malaysian Borneo), An. dirus (in Vietnam)12 and An. hackeri and

An. cracens (in Peninsular Malaysia)1,13.

People that are at risk of acquiring knowlesi malaria are those that

enter thehabitat of themacaque reservoir hosts and theAnopheline

vectors at dusk or later as this coincides with the peak biting time of

the vectors14,15. These include subsistence farmers, timber camp

workers, hunters, army personnel and also travelers to forests or

forest-fringe areas. Visitors to South-east Asia from Australia, USA,

Finland, Sweden,Germany, France, NewZealand, Taiwan and Japan

have acquired knowlesi malaria following holidays or working visits

Under theMicroscope


to Malaysian Borneo, Peninsular Malaysia, Brunei, Thailand,

Indonesia and the Philippines16.

Molecular and whole genome studies

In order to understand the molecular epidemiology and demo-

graphic history of knowlesimalaria, themitochondrial (mt) genome

sequences of P. knowlesi were initially studied17. Certain mt hap-

lotypes were shared between humans and macaques and there

were no haplotypes that were associated exclusively with either

host; further evidence supportingP.knowlesi as a zoonoticparasite.

Additional analyses indicated that P. knowlesi was as old as, if not

older than, P. falciparum and P. vivax, and that it underwent a

population expansion between 30,000 to 40,000 years ago. Maca-

ques colonized Asia over 5million years ago18 and are probably the

original hosts forP. knowlesi. A recent study, where 599P. knowlesi

samples from Peninsular Malaysia and Malaysian Borneo were

analysed by a panel of ten microsatellite markers, showed there

are twohighly divergent sub-populations ofP. knowlesi, andeachof

these subpopulations correspond with parasites from either long-

tailed or pig-tailed macaques19. More recently, genome-wide se-

quence analysis of clinical P. knowlesi isolates from Malaysian

Borneo shows sub-population structure that matches the analysis

using microsatellite markers and also demonstrate there is a third

sub-population of parasites, corresponding to laboratory strains

isolated over 50 years ago from Peninsular Malaysia and the

Philippines20. No signals of positive selection were observed in

P. knowlesi around five orthologues of known P. falciparum

drug resistance genes, indicating that the parasites in the reservoir

macaque hosts have not been under antimalarial drug selection,

thereby providing further evidence that knowlesi malaria is a

zoonosis.

Diagnosis

In laboratories in malaria-endemic countries, malaria is diagnosed

by examination of blood films by microscopy. Under the micro-

scope, the early blood forms of P. knowlesi are identical to those

of P. falciparum, while the other developmental stages, including

the ‘band forms’, are similar to those of P. malariae3. There are

minor morphological differences between these two species. The

mature schizonts of P. knowlesi can contain up to 16 merozoites,

whereas those of P. malariae have between 6–123. However,

mature schizonts are not found in all blood films examined and in

diagnostic laboratories, where technologists are only trained to

recognise P. falciparum, P. vivax, P. ovale and P. malariae, most

P. knowlesi infections have been identified by microscopy as

P. malariae2,4,21. Although morphologically similar, P. knowlesi

parasites multiply every 24 h in the blood while this erythrocytic

cycle is 72 h for P. malariae9.

Molecular detection methods are the most sensitive and accurate

techniques for identification of P. knowlesi. These include single

and nested PCR assays, real-time PCR assays and loop-mediated

isothermal assays4. However, these assays are relatively expensive,

not rapid and are not readily available in resource-poor laboratories

where the majority of P. knowlesi infections are detected. Rapid

diagnostic tests (RDTs) for malaria are available, but the overall

sensitivity of detection of a small number of RDTs that have been

evaluated against knowlesi malaria cases varied between 26–74%

andwas even lower (0–45%) for parasitaemias below1000parasites/

mL22–24. Due to the rapid multiplication rate of P. knowlesi in

the blood of 24 h, sensitive RDTs capable of detecting knowlesi

malaria at the early phase of infection are urgently required for rural

laboratories.

Clinical and laboratory features

of knowlesi malaria

P. knowlesi causes a wide spectrum of disease, from asymptomatic

infections9,25 to fatal ones26–29. Themost common presenting signs

and symptoms reported are fever with chills, followed by headache,

myalgia, poor appetite, arthralgia, cough, abdominal pain and

diarrhoea27. These are not significantly different to those observed

in patients with vivax and falciparum malaria. The majority of cases

(93.5%27 and 84.5%30) at district hospitals in Sarawak had uncom-

plicated malaria with a fatality rate of 2%, whereas in a retrospective

study in a referral hospital in Sabah, 61% of 56 cases were uncom-

plicated and the fatality rate was 27%31. However, subsequently at

the same referral hospital, the use of intravenous artesunate for

severe malaria cases and artemisinin combination therapy for non-

severe cases, resulted in no deaths among 130 knowlesi malaria

patients29. Typical complications of severe knowlesi malaria in

adults include jaundice, acute kidney injury, hypotension, acute

respiratory distress syndrome andmetabolic acidosis26,27,29,30,32. In

adults, severe anaemia has not been observed and neither has

cerebral malaria, while severe disease has not been noted in the

relatively small number of children with knowlesi malaria4,33.

Thrombocytopaenia is very common, occurring in 97.3 to 100% of

knowlesi malaria patients, and together with parasitaemia, corre-

lates with severity of disease27,30,31. Following a case control study,

it was recommended that any patient with a platelet count of

<45 000/mL or parasitaemia of >35 000 parasites/mL should be

regarded at risk of developing complications and should be treated

for severe malaria30.

Under theMicroscope


Treatment of knowlesi malaria

Since knowlesi malaria is primarily a zoonosis, the parasites have

beenunder no antimalarial drug pressure and should be susceptible

to all antimalarials. This has been observed in hospital-based studies

as well as case reports where several antimalarials have been used

successfully to treat knowlesi malaria patients4. P. knowlesi para-

sites are highly sensitive to chloroquine34 but following an informal

consultation on the public health importance of knowlesi malaria

organised by the WHO in 2011, it was recommended that in areas

where knowlesi malaria has been detected, all infections diagnosed

as P. malariae by microscopy should be treated and managed as

for falciparum malaria35. Therefore, for uncomplicated knowlesi

malaria cases in South-east Asia, artemisinin combination therapy is

recommended. For severe knowlesi malaria, intravenous antimalar-

ials should be administered and the use of artesunate in a tertiary

referral hospital in Sabah was associated with zero mortality29.

Future directions

The available molecular, entomological and epidemiological data

strongly indicate that knowlesi malaria is primarily a zoonosis.

However, human-to-human transmission has been demonstrated

under experimental conditions9 and it is not known whether it is

currently occurring. The reasons for the increase in the number of

knowlesi malaria cases, particularly in Malaysian Borneo, are also

unknown. Whether the increase is due to increased awareness,

changes in the feeding habits of the vectors, the destruction of the

natural habitats of themacaque reservoir, humanmigration to areas

close to macaque habitats, a recent adaptation of knowlesi malaria

parasites to humans, or to some other factors needs to be investi-

gated. In addition, currently available methods of control of human

malaria involving the use of insecticide treated bednets and residual

spraying of houses are ineffective against knowlesi malaria, where

transmission primarily occurs outdoors. Therefore, effective meth-

odsofpreventionandcontrolneed tobe foundand implemented, in

order to prevent P. knowlesi from establishing itself in the human

population.

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Pathog. 11, e1004888. doi:10.1371/journal.ppat.1004888

20. Assefa, S. et al. (2015) Population genomic structure and adaptation in the

zoonotic malaria parasite Plasmodium knowlesi. Proc. Natl. Acad. Sci. USA

112, 13027–13032. doi:10.1073/pnas.1509534112

21. William, T. et al. (2014) Changing epidemiology of malaria in Sabah, Malaysia:

increasing incidence of Plasmodium knowlesi. Malar. J. 13, 390. doi:10.1186/

1475-2875-13-390

22. Foster, D. et al. (2014) Evaluation of three rapid diagnostic tests for the detection

of human infections with Plasmodium knowlesi. Malar. J. 13, 60. doi:10.1186/

1475-2875-13-60

23. Barber, B.E. et al. (2013) Evaluation of the sensitivity of a pLDH-based and an

aldolase-based rapid diagnostic test for diagnosis of uncomplicated and severe

malaria caused by PCR-confirmed Plasmodium knowlesi, Plasmodium falci-

parum, and Plasmodium vivax. J. Clin. Microbiol. 51, 1118–1123. doi:10.1128/

JCM.03285-12

24. Grigg, M.J. et al. (2014) Combining parasite lactate dehydrogenase-based and

histidine-rich protein 2-based rapid tests to improve specificity for diagnosis of

malaria due to Plasmodium knowlesi and other Plasmodium species in Sabah,

Malaysia. J. Clin. Microbiol. 52, 2053–2060. doi:10.1128/JCM.00181-14

25. Fornace, K.M. et al. (2015) Asymptomatic and submicroscopic carriage of

Plasmodium knowlesi malaria in household and community members of

clinical cases in Sabah, Malaysia. J. Infect. Dis. doi:10.1093/infdis/jiv475

Under theMicroscope


26. Cox-Singh, J. et al. (2008) Plasmodium knowlesi malaria in humans is widely

distributed and potentially life threatening. Clin. Infect. Dis. 46, 165–171.

doi:10.1086/524888

27. Daneshvar, C. et al. (2009) Clinical and laboratory features of human

Plasmodium knowlesi infection. Clin. Infect. Dis. 49, 852–860. doi:10.1086/

605439

28. Rajahram,G.S.et al. (2012)Deathsdue toPlasmodiumknowlesimalaria in Sabah,

Malaysia: association with reporting as Plasmodium malariae and delayed

parenteral artesunate. Malar. J. 11, 284. doi:10.1186/1475-2875-11-284

29. Barber, B.E. et al. (2013) A prospective comparative study of knowlesi,

falciparum, and vivax malaria in Sabah, Malaysia: high proportion with severe

disease from Plasmodium knowlesi and Plasmodium vivax but no mortality

with early referral and artesunate therapy. Clin. Infect. Dis. 56, 383–397.

doi:10.1093/cid/cis902

30. Willmann, M. et al. (2012) Laboratory markers of disease severity in Plasmodium

knowlesi infection: a case control study. Malar. J. 11, 363. doi:10.1186/1475-

2875-11-363

31. William, T. et al. (2011) Severe Plasmodium knowlesi malaria in a tertiary care

hospital, Sabah, Malaysia. Emerg. Infect. Dis. 17, 1248–1255. doi:10.3201/eid

1707.101017

32. Cox-Singh, J. et al. (2010) Severe malaria – a case of fatal Plasmodium knowlesi

infection with post-mortem findings: a case report. Malar. J. 9, 10. doi:10.1186/

1475-2875-9-10

33. Barber, B.E. et al. (2011)Plasmodiumknowlesimalaria in children.Emerg. Infect.

Dis. 17, 814–820. doi:10.3201/eid1705.101489

34. Daneshvar, C. et al. (2010) Clinical and parasitological response to oral chloro-

quine and primaquine in uncomplicated human Plasmodium knowlesi infec-

tions. Malar. J. 9, 238. doi:10.1186/1475-2875-9-238

35. WHO (2011) Informal consultation on the public health importance of Plasmo-

diumknowlesi.WorldHealthOrganizationRegionalOffice for theWesternPacific

Press.

Biography

Professor Balbir Singh is the Director of the Malaria Research

Centre at University Malaysia Sarawak. His research interests

include the epidemiology, pathogenesis and evolution of malaria

parasites.

Under theMicroscope


Diagnosis of human taeniasis

Abdul Jabbar, Charles Gauci and Marshall W Lightowlers

Faculty of Veterinary and Agricultural Sciences, The University of Melbourne, Werribee, Vic. 3030, Australia, Email: [email protected]

Taenia solium, T. saginata and T. asiatica are taeniid tape-

worms that cause taeniasis in humans and cysticercosis in

intermediate host animals. T. solium can also cause cysti-

cercosis in humans. A number of diagnostic methods have

been developed to diagnose Taenia species that infect

humans. This article is aimed at providing an overview of

currently available diagnosticmethods for human taeniasis.

Human taeniasis is an important zoonotic health problem and is

causedby adult stages of taeniid cestodes, includingTaenia solium,

T. saginata, and T. asiatica. These parasites have indirect life cycles

wherehumansact as definitivehostswhereaspigs (forT. solium and

T. asiatica) and cattle (T. saginata) serve as intermediate hosts.

Humans become infected by ingesting parasite cysts present in raw

or undercooked infected meat/liver; when the cyst reaches the

human intestine, it develops into an adult tapeworm, releasing

segments and/or eggs in the stools or motile segments (e.g.

T. saginata) are expelled actively. Intermediate hosts such as cattle

and pigs become infected when they ingest taeniid eggs via con-

taminated feed or water and the larval stage (cysticercus) forms in

muscular and sometimes other tissues1. The adult tapeworm stage

of Taenia spp. is relatively innocuous and does not cause patho-

genic effects in humans; however, the intermediate stage of

T. solium can also develop in human brains causing neurocysticer-

cosis, a major cause of neurological disease in many developing

countries as well as other organs causing intramuscular, ocular,

subcutaneous and spinal cysticercoses2,3. T. saginata is endemic in

Australia, whereas T. solium and T. asiatica are exotic. However,

people coming and/or returning to Australia from endemic coun-

tries can be infected with T. solium, leading to the possibility that

infected individualsmaypass segmentsof theparasite in their stools,

which can serve as a source of infection for human cysticercosis.

Therefore, differentiation ofTaenia species becomes significant for

surveillance and control of human taeniasis.

A number of diagnosticmethods have beenused todifferentiate the

common human cestodes, T. saginata and T. solium; however,

each method has its advantages and disadvantages, and careful

attention should be paid to determining which particular test is

best to use for differentiation of the two species4. The following

sections provide a quick rundown on various diagnostic methods

available to differentiate Taenia spp. that infect humans.

Microscopic diagnosisTraditionally, diagnosis of taeniasis has been based on the detection

of eggs by microscopic examination but this method lacks

sensitivity and specificity as T. solium and T. saginata eggs are

morphologically identical, making species identification impossi-

ble5.However,morphological examinationof gravidproglottids can

allow the differentiation of T. solium and T. saginata provided the

internal structures (i.e. uterine branches) are intact5. Sometimes,

even the morphological examination of proglottids does not allow

the differentiation of T. solium and T. saginata, thus requiring

alternate methods for the differentiation of human Taenia spp.

ImmunodiagnosisThefirstmethodof coproantigen (parasite antigens inhuman stool)

detection using enzyme-linked immunosorbent assay (ELISA) was

developed by Allan et al.6. Although the test displayed a higher

sensitivity and specificity than microscopic diagnosis for the detec-

tion of Taenia spp.7, it did not allow differentiation of T. saginata

andT. solium. Recently,Guezala et al.8 developed another coproan-

tigen ELISA and successfully differentiated T. solium from

T. saginata. These tests were developed in individual labs using

Under theMicroscope


in-house reagents and have not been independently validated in

different laboratories nor have they been widely used in diagnostic

laboratories.

The first serological assay to detect specific antibodies against

T. solium infection in humans was developed by Wilkins et al.9

Subsequently, a number of studies reported various immunoassays

for the diagnosis of human taeniasis, primarily caused by

T. solium10,11 and these studies used either native excretory-secre-

tory products collected from adult tapeworms9 or cloned and

expressed excretory-secretory products of adult T. solium10,11. Like

coproantigen ELISAs, the detection of Taenia species-specific

antibodies are also more specific and sensitive than microscopic

techniques. However, these tests have been found to have some

degree of cross-reactivity in sera from patients with cystic echino-

coccosis, ascariasis, and schistosomiasis11. Furthermore, currently

available immunodiagnostic tests may give false positive results

as specific circulating antibodies in taeniasis patients could possibly

remain detectable for some time either after treatment and recent

past infections4.

Molecular diagnosisA number of molecular methods using PCR-based technologies

have been developed to either determine the presence of Taenia

species-specific DNA in human stools or differentiate Taenia

spp. (T. solium, T. saginata or T. asiatica) based on the analysis

of DNA extracted directly from tapeworm12. PCR-based methods

have higher sensitivity in the detection of taeniasis cases (i.e. the

detection of parasite DNA in human stools) than microscopy

alone. In addition, a combined use of PCR and microscopy have

been found to improve diagnostic sensitivity where Yamasaki

et al.13 showed that some proven egg-positive cases were negative

by PCR. Specificity of PCR is high with control faecal samples,

including samples from patients with other parasitic infections,

being almost always negative in PCR.

To date, predominantly conventional and multiplex PCR, and PCR-

restriction fragment lengthpolymorphism (PCR-RFLP) haveutilized

various markers, including internal transcribed spacer, mitochon-

drial cytochrome c oxidase subunit I gene as well as 12S rDNA,

cathepsin L-like cysteine peptidase, T. saginata-specific repetitive

sequence (HDP1) and cestode-specific sequence (HDP2) to dis-

criminate between human Taenia spp13–21 (Table 1). Mayta et al.19

developed a nested PCRutilizing two rounds of PCR amplification of

the Tso31 gene, which is more sensitive than conventional PCR but

pose technical difficulties. A field DNA-based test known as loop

mediated isothermal amplification (LAMP) was developed by

Nkouawa et al.20, which amplifies the cox1 gene. To date, only one

real-time PCR to discriminate T. solium and T. saginata has been

developed which targets the internal transcribed spacer 1 of

the nuclear ribosomal RNA21. The majority of DNA-based methods

have utilised DNA isolated from proglottids for Taenia species

identification while relatively fewer studies isolated DNA from

stool samples (Table 1). In addition, almost all studies have been

testedonlyon small numbersof usually knownpositive andnegative

samples.

Conclusions and future perspectivesHuman taeniasis is a worldwide parasitic disease and detection and

discrimination of T. solium, T. saginata and T. asiatica remains a

Table 1. Selected PCR-based assays for detection and characterization of human Taenia spp.

Taenia species Molecular method Target(s) Sample type(s) Reference

T. saginata PCR Gel HDP1 Proglottids 14

T. solium, T. saginata Multiplex PCR HDP2 Proglottids 15

T. solium, T. saginata Multiplex PCR-RFLP HDP2 Proglottids 16

T. solium, T. saginata,T. asiatica

PCR-RFLP Mitochondrial 12S rDNA Proglottids 17


Multiplex PCR cox1 Proglottids, stool 13

T. solium, T. saginata PCR-RFLP cox1 Stool 18

T. solium Nested PCR Tsol31 Stool 19


LAMP Clp, cox1 Proglottids, cysticerci, stool 20

T. solium, T. saginata Real-time multiplex PCR ITS1 Stool 21

RFLP, restriction fragment length polymorphism; ITS, internal transcribed spacer; cox1,mitochondrial cytochrome coxidase subunit I gene; Clp, cathepsin L-likecysteine peptidase; HDP1, T. saginata-specific repetitive sequence; HDP2, cestode-specific sequence. Adapted from Verweij and Stensvold12.

Under theMicroscope


public health concern. Microscopic, immunological and molecular

methods have been used to detect and differentiate Taenia spp.,

and a combination of two or more methods appears to provide

higher sensitivity13,21. Almost all of the immunological and molec-

ular methods developed so far have not been independently tested

and validated on controlled negative and positive samples as well as

field samples. Amongexistingmolecularmethods, thenestedPCR19

provides thehighest sensitivity and specificityof thosemethods that

have been developed and validated using unselected faecal samples

fromparasitological proven taeniasis carriers. Although themethod

is technically challenging and could be expected to be relatively

expensive to use, the reagents required are available worldwide

and laboratories capable of undertaking PCR competently can run

this PCR4.

To the best of our knowledge, no standardised PCRs are available in

Australia to discriminate human Taenia spp. Future studies should

focus on independent validation of the nested PCR19 and the LAMP

technique20 as these two DNA-based tests offer higher sensitivity

and a user-friendly option without sophisticated equipment, re-

spectively. In addition, future studies should be aimed at extracting

DNA from sodium acetate-acetic acid-formalin (SAF) fixed proglot-

tidsusingdifferentmethods as thecurrentDNAextractionprotocols

do not provide reliable and consistent DNA yield for PCRs (Jabbar,

unpublished data). To date only one study has reported the extrac-

tion of DNA from long-term stored Taenia specimens in formalin22

and this has not been independently verified.

References1. Roberts, L.S. et al. (2013) Foundations of Parasitology, 9th edn. McGraw-Hill

Education, USA.

2. Mahanty, S. et al. (2010) Cysticercosis and neurocysticercosis as pathogens

affecting the nervous system. Prog. Neurobiol. 91, 172–184. doi:10.1016/

j.pneurobio.2009.12.008

3. García, H.H. et al. (2003) Taenia solium cysticercosis. Lancet 362, 547–556.

doi:10.1016/S0140-6736(03)14117-7

4. Lightowlers, M.W. et al. (2015) Monitoring the outcomes of interventions against

Taenia solium: option and suggestion. Parasite Immunol. In press. doi:10.1111/

pim.12291

5. Chapman, A. et al. (1995) Isolation and characterization of species-specific

DNA probes from Taenia solium and Taenia saginata and their use in an egg

detection assay. J. Clin. Microbiol. 99, 1283–1288.

6. Allan, J.C. et al. (1990) Immunodiagnosis of taeniasis by coproantigen detection.

Parasitology 101, 473–477. doi:10.1017/S0031182000060686

7. Allan, J.C. et al. (1996) Field trial of the coproantigen-based diagnosis of Taenia

solium taeniasis by enzymelinked immunosorbent assay. Am. J. Trop. Med. Hyg.

54, 352–356.

8. Guezala, M.C. et al. (2009) Development of a species-specific coproantigen ELISA

for human Taenia solium taeniasis. Am. J. Trop. Med. Hyg. 81, 433–437.

9. Wilkins, P.P. et al. (1999) Development of a serologic assay to detect Taenia

solium taeniasis. Am. J. Trop. Med. Hyg. 60, 199–204.

10. Levine, M.Z. et al. (2004) Characterization, cloning, and expression of two

diagnostic antigens for Taenia solium tapeworm infection. J. Parasitol. 90,

631–638. doi:10.1645/GE-189R

11. Levine, M.Z. et al. (2007) Development of an enzyme-linked immunoelectro-

transfer blot (EITB) assay using two baculovirus expressed recombinant antigens

for diagnosis of Taenia solium taeniasis. J. Parasitol. 93, 409–417. doi:10.1645/

GE-938R.1

12. Verweij, J.J. and Stensvold, C.R. (2014) Molecular testing for clinical diagnosis and

epidemiological investigations of intestinal parasitic infections. Clin. Microbiol.

Rev. 27, 371–418. doi:10.1128/CMR.00122-13

13. Yamasaki, H. et al. (2003) Cysticercosis/taeniasis: recent advances in serological

andmolecular diagnoses. Southeast Asian J. Trop.Med. PublicHealth34, 98–102.

14. Gonzalez, L.M. (2000) Differential diagnosis of Taenia saginata and Taenia

solium infection by PCR. J. Clin. Microbiol. 38, 737–744.

15. González, L.M. et al. (2002) Differential diagnosis of Taenia saginata and Taenia

solium infections: from DNA probes to polymerase chain reaction. Trans. R. Soc.

Trop. Med. Hyg. 96, S243–S250. doi:10.1016/S0035-9203(02)90083-0

16. González, L.M. et al. (2002) PCR tools for the differential diagnosis of Taenia

saginata and Taenia solium taeniasis/cysticercosis from different geographical

locations.Diagn.Microbiol. Infect. Dis.42, 243–249. doi:10.1016/S0732-8893(01)

00356-X

17. Rodriguez-Hidalgo, R. et al. (2002) Comparison of conventional techniques to

differentiate between Taenia solium and Taenia saginata and an improved

polymerase chain reaction-restriction fragment length polymorphism assay using

a mitochondrial 12S rDNA fragment. J. Parasitol. 88, 1007–1011. doi:10.1645/

0022-3395(2002)088[1007:COCTTD]2.0.CO;2

18. Nunes, C.M. et al. (2005) Taenia saginata: differential diagnosis of human

taeniasis by polymerase chain reaction-restriction fragment length polymorphism

assay. Exp. Parasitol. 110, 412–415. doi:10.1016/j.exppara.2005.03.008

19. Mayta, H. et al. (2008) Nested PCR for specific diagnosis of Taenia solium

taeniasis. J. Clin. Microbiol. 46, 286–289. doi:10.1128/JCM.01172-07

20. Nkouawa, A. et al. (2009) Loop-mediated isothermal amplification method for

differentiation and rapid detection of Taenia species. J. Clin. Microbiol. 47,

168–174. doi:10.1128/JCM.01573-08

21. Praet, N. et al. (2013) Bayesian modelling to estimate the test characteristics of

coprology, coproantigen ELISA and a novel real-time PCR for the diagnosis of

taeniasis. Trop. Med. Int. Health 18, 608–614. doi:10.1111/tmi.12089

22. Jeon, H.-K. et al. (2011) Molecular identification of Taenia specimens after long-

term preservation in formalin. Parasitol. Int. 60, 203–205. doi:10.1016/j.parint.

2010.12.001

BiographiesDr Abdul Jabbar is a Senior Lecturer in Veterinary Parasitology at

The University of Melbourne. His main research interests cover

epidemiology and diagnosis of parasites of socioeconomic impor-

tance using next-generation molecular tools.

Dr Charles Gauci is a Senior Research Fellow at The University of

Melbourne. He has worked with Prof Marshall Lightowlers through-

out his career and his research interests focus on recombinant

vaccines for prevention of transmission of the parasite causing

neurocysticercosis and the related parasite that causes hydatid

disease.

Professor Marshall W Lightowlers has been a full-time research

scientist supported bymedical research funding formore-or-less all

of his working life. He currently holds appointments as Laureate

Professor at the University of Melbourne’s Faculty of Veterinary

and Agricultural Sciences, and Principal Research Fellow with the

NHMRC.

Under theMicroscope


Protozoa PCR: boon or bane

Colin Pham

St Vincent’s HospitalMelbourne, Vic., AustraliaEmail: [email protected]

Harsha Sheorey

St Vincent’s HospitalMelbourne, Vic., AustraliaEmail: [email protected]

Parasitedetection in faeceshas traditionallybeenperformed

by microscopy, a procedure that is labour-intensive and

highly specialised. In addition, identification bymicroscopy

based on morphological features alone is subjective and

prone to wide variability. Although enzyme immunoassays

(EIA) of high sensitivity have been developed1 they can

detect only a limited range of pathogens. Given these

factors the introduction of Polymerase Chain Reaction

(PCR) into the routine diagnostic laboratory has improved

parasite detection rates2. The ability to multiplex has en-

abled the detection of multiple targets from a single sample

and provides an objective alternative to identification by

morphology.

Results

In February 2014, St Vincent’s Pathology, Melbourne, introduced a

new testing algorithm making use of a commercial multiplex PCR

(LightMix�Gastroenteritis Parasite Kit, Roche Molecular Systems,

Pleasanton, California, USA) for parasite detection. This assay is

designed to detect Giardia intestinalis, Cryptosporidium species,

Dientamoeba fragilis and Entamoeba histolytica, four human gas-

trointestinal protozoanpathogens3.Blastocystiswasnot includedasa

target in thismultiplex assay, as theprimersdonotdifferentiate the17

differentsubtypesnowknown4,manybeingofanimaloriginandmost

of doubtful clinical significance.

Since the commencement of testing by PCR, the rates of detection

have improved when compared with results obtained based on

the previous protocol that included microscopy and Giardia/

Cryptosporidium antigen testing (Table 1). Comparison over a

three month period showed that the range of common parasites

detected in the population remains similar with D. fragilis

accounting for the majority of the additional parasites detected

(Figure 1). The reduction of Cryptosporidium cases from 18 (0.8%)

to 4 (0.4%) was not statistically significant and is thought to relate

to seasonal variation in prevalence. The additional E. histolytica

cases were detected from microscopy-positive specimens referred

from other laboratories. In addition, the assay was positive for

Table 1. Three months comparison between pre and post implementation of protozoa PCR testing.

Quarter 2: 2013Pre PCR

Quarter 2: 2014Post PCR

Statistical significance

Samples 2198 1096 P < 0.001

G. intestinalis 24 (1.1%) 33 (3.0%) P < 0.001

C. parvum 18 (0.8%) 4 (0.4%) P= 0.132

E. histolytica – 4 (0.4%) P= 0.005

D. fragilis 47 (2.1%) 163 (14.9%) P < 0.001

Total positives 89 (4%) 204 (19%) P < 0.001

LabReport


E. histolytica in liver aspirates from four cases of suspected amoebic

hepatitis.

The detection of protozoa by molecular technology at St Vincent’s

Pathology, Melbourne, over the 15 month period since implemen-

tation of the multiplex PCR assay is shown in Table 2. The rate of

D. fragilis detection increased from 2.1% by microscopy to 15.3%

with PCR; this represents the most significant finding, accounting

for 78% of all parasites detected by this test.

Discussion

The advent of PCRhas definitely become a ‘boon’ for our laboratory

in providing a rapid, less labour-intensive method for detecting the

threepathogenicprotozoaG. intestinalis,Cryptosporidium species

and E. histolytica. PCR can detect low numbers easily missed by

microscopy and also candifferentiate pathogenicE. histolytica from

morphologically similar, non-pathogenic Enta-moeba dispar/

moshkovskii5. During the initial validation period, 10 Entamoeba

species identified by microscopy were tested by PCR and only two

of these samples were positive for E. histolytica. The ability to

differentiate betweenpathogenic E. histolytica andnon-pathogenic

Entamoeba species is clinically significant. Results concur with a

prevalence study of Entamoeba in immigrants in Spain that showed

that the majority of cyst-positive samples detected by microscopy

were E. dispar and do not require treatment5.

On the other hand, PCR seemed to become a ‘bane’ by detecting a

large number of D. fragilis in asymptomatic people, especially

children6,7. Numerous parents of ‘well’ children get anxious about

this so called ‘bad bug that does not go away’. Many of these

‘worried well’ get treated with various antiparasitic drugs and

combinations (some of which are known to imbalance the normal

gut flora). They ‘doctor shop’ and are being referred to specialists

such as paediatricians, gastroenterologists and infectious diseases

physicians (communications with General Practitioners and

Australia and New Zealand Paediatric Infectious Diseases Group).

The reliance on morphological diagnosis by microscopy of fixed

stained-smears and the recognition that parasite shedding may be

intermittent, suggests that prior to PCR diagnosis, the true preva-

lence of this protozoon has been underestimated. A review of

D. fragilis carriage has shown that prevalence ranges from 0.2% to

82% in numerous geographical settings, utilising a variety of detec-

tion methods8. In this review, the authors conclude that in

‘symptomatic patients who harbor Dientamoeba and no other

pathogen, it should be considered as the etiological agent and

treated’. However, they also mention that evidence is only circum-

stantial. There has been debate about the clinical significance of

D. fragilisgivensuchwidevariation inprevalenceestimates coupled

with theobservation that theprotozoamaybe found in symptomatic

and asymptomatic individuals and variable responses to antipara-

sitic drugs. A recent case-controlled study by Krogsgaard et al.6

demonstrated that D. fragilis and Blastocystis were detected in a

greater proportion of faecal samples from the asymptomatic back-

ground population in Denmark than from subjects with irritable

bowel syndrome. Röser et al.7 presented data from four years of

routine PCR testing and found that from a total of 22,484 samples,

43% were positive for D. fragilis. With the introduction of PCR

into routine testing within our laboratory, our data suggest that

the prevalence of asymptomatic D. fragilis carriage is higher than

previously reported.

Of all D. fragilis detected in our study period, 40% of faeces were

formed, 58% were semi-formed or unformed and only 2% were

loose/liquid as observed in the laboratory. The clinical notes pro-

vided on pathology request forms ranged from diarrhoea to vague

0

20

40

60

80

100

120

140

160

180

Quarter 2 – 2013: Pre PCR Quarter 2 – 2014: Post PCR

Tot

al p

ositi

ves

G. intestinalis

Cryptosporidium spp.

E. histolytica

D. fragilis

Figure 1. Three months comparison of positive parasite cases pre andpost implementation of protozoa PCR.

Table 2. Protozoal infections detected by multiplex PCR February2014 to May 2015.

Total positive(percentage)

% of allpositives

D. fragilis 828 (15.3%) 78.0

G. intestinalis 183 (3.4%) 17.2

Cryptosporidiumspp.

34 (0.6%) 3.2

E. histolyticaA 13 (0.24%) 1.22

Total positives 1062 (19.7%) 100

Total tested 5384 –

AAdditional four liver aspirates were positive by PCR.

LabReport


IBS like symptoms such as abdominal pain or discomfort, altered

bowel habit and others unrelated to gastrointestinal tract. Samples

that were positive for D. fragilis by PCR were predominantly in

children below 10 years with a second peak in young adults in age

group 30–40 years (Figure 2), a similar distribution to that described

by Röser et al.7.

With the more widespread adoption of faecal parasite multiplex

PCR assays, the higher positive rate of D. fragilis by PCR reported

by laboratories (especially when diagnosed in those with non-

specific symptoms), is confusing and of concern to clinicians. Many

patients get unnecessary courses of antiparasitic drugs with more

than half not clearing the parasite. A double-blinded, placebo-

controlled metronidazole study by Röser et al.9 suggests that

treatment is not associated with better clinical outcomes. Apart

from optimal therapy not being known and a highly variable re-

sponse to treatment, the pathogenicity of D. fragilis has not been

reliably demonstrated.

It must be noted that the use of molecular technology is limited to

the targeted screening of only a relatively small number of parasites

in a low prevalence population. Microscopy still plays a key role in

less common parasite identification, especially in specific patient

groups where parasites other than those targeted by this multiplex

are suspected. Forparasite examination inpopulations fromregions

where there is high parasite endemicity, microscopy remains the

‘gold standard’ with the capacity to screen for the majority of

pathogens.

In summary, molecular testing can offer accurate results for the

detection of the four commonly seen parasites in low prevalence

populations. It has been shown to be comparable to EIA for

G. intestinalis and Cryptosporidium diagnosis and superior to

microscopy for D. fragilis and E. histolytica detection. Our new

testing algorithm for parasite detection utilising PCR has shown to

improve sensitivity and specificity. It has significantly reduced the

time taken to stainfixed smears andmicroscopy, allowing for amore

efficient use of labour, whilst improving parasite detection rates.

However, laboratories should consider the limitations of reporting

parasites of doubtful clinical significance such as D. fragilis and

Blastocystis species, and should alert physicians of the possibility of

this being non-pathogenic in immunocompetent people.

References1. Vidal, A.M. andCatapani,W.R. (2005) Enzyme-linked immunosorbent assay (ELISA)

immunoassaying versus microscopy: advantages and drawbacks for diagnosing

giardiasis. SaoPauloMed. J.123, 282–285.doi:10.1590/S1516-31802005000600006

2. Bruijnesteijn van Coppenraet, L.E. et al. (2009) Parasitological diagnosis combining

an internally controlled real-time PCR assay for the detection of four protozoa in

stool samples with a testing algorithm for microscopy. Clin. Microbiol. Infect. 15,

869–874. doi:10.1111/j.1469-0691.2009.02894.x

3. Slack, A. (2012) Parasitic causes of prolonged diarrhoea in travellers – diagnosis and

management. Aust. Fam. Physician 41, 782–786.

0

5

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1 11 21 31 41 51 61 71 81

Tot

al D

. fra

gilis

pos

itive

s

Age

Figure 2. Age distribution of patients positive for D. fragilis by PCR in our study.

Summary of current faecal protozoa molecular testing:

* Very good at detecting low numbers of known protozoan parasites such asE. histolytica, Cryptosporidium species and Giardia intestinalis

* Useful for discriminating between invasive E. histolytica andnon-pathogenic Entamoeba species

* Detects protozoa with doubtful pathogenicity such as D. fragilis at a higherrate when compared to microscopy, leading to confusion, anxiety andunnecessary treatment

* Cannot differentiate animal subtypes of Blastocystis (non-pathogenic) fromhuman subtypes (of debatable pathogenicity)

LabReport


4. Stensvold, C.R. (2013) Blastocystis: genetic diversity and molecular methods for

diagnosis and epidemiology. Trop. Parasitol. 3, 26–34. doi:10.4103/2229-5070.

113896

5. Gutiérrez-Cisneros, M.J. et al. (2010) Application of real-time PCR for the differ-

entiation of Entamoeba histolytica and E. dispar in cyst-positive faecal samples

from 130 immigrants living in Spain. Ann. Trop. Med. Parasitol. 104, 145–149.

doi:10.1179/136485910X12607012373759

6. Krogsgaard, L.R. et al. (2015) The prevalence of intestinal parasites is not greater

among individuals with irritable bowel syndrome: a population-based case-control

study. Clin. Gastroenterol. Hepatol. 13, 507–513.

7. Röser, D. et al. (2013) Dientamoeba fragilis in Denmark: epidemiological expe-

rience derived from four years of routine real-time PCR. Eur. J. Clin. Microbiol.

Infect. Dis. 32, 1303–1310. doi:10.1007/s10096-013-1880-2

8. Barratt, J.L. et al. (2011) A review of Dientamoeba fragilis carriage in humans:

several reasons why this organism should be considered in the diagnosis of

gastrointestinal illness. Gut Microbes 2, 3–12. doi:10.4161/gmic.2.1.14755

9. Röser, D. et al. (2014) Metronidazole therapy for treating dientamoebiasis in

children is not associated with better clinical outcomes: a randomized, double-

blinded and placebo-controlled clinical trial. Clin. Infect. Dis. 58, 1692–1699.

doi:10.1093/cid/ciu188

Biographies

Colin Pham, is a Medical Scientist with over 10 years’ experience

in bacteriology, serology and molecular testing. He underwent his

training at St Vincent’s Melbourne in 2003, and started his career

in Microbiology at the Monash Medical Centre. Since re-joining

St Vincent’s, he has specialised in molecular testing and emerging

molecular technology. He completed a Masters of Applied Science

(Medical Science) at RMIT in 2014, with his thesis titled ‘Impact

of molecular testing on the laboratory diagnosis of Giardia

intestinalis, Cryptosporidium parvum, Dientamoeba fragilis and

Entamoeba histolytica from clinical stool samples’.

Dr Harsha Sheorey, is a Clinical Microbiologist at St Vincent’s

Hospital in Melbourne. His special interest is in clinical parasitology

and tropical medicine and is the author of Clinical Parasitology –

a handbook for medical practitioners and microbiologists: a

second edition has recently been published. He is an invited writer

for the Nematodes chapter in ASMManual of Clinical Microbiology.

He coordinates the Victorian branch of Parasitology & Tropical

Medicine SIG and is actively involved in organisation of the Austra-

lian Parasitology and Tropical Medicine Master Class.

LabReport


FT020 Water Microbiology Australian StandardsCommitteeRobin Woodward

Email: [email protected]

I am the Australian Society of Microbiology representative on the

FT020 Water Microbiology Australian Standards Committee. This

committee meets when there are Australian Standards (AS) that are

up for review (approximately every 5 years) or when a new Standard

has been proposed and requires work. Before I became a member

on this committee I thought AS were put together by an elite group

of people and that was all they did. The Committee is made up of

elite members, (of course that goes without saying �) but we are

peoplewhowork in the industry andmost stillwork in the laboratory

or have at some time in our career. There is the Standards Australia

secretarywho ensures themeeting and Standards follow the correct

protocols, the Chair who runs the meeting, a NATA representative

and members comprising people like myself who are in a supervi-

sory role but still perform bench work in the real world. I still do

bench work and that is why I enjoy the challenge of being on

the Committee. It entails reading all the Standards, assessing their

relevance and at meetings discussing any issues that may arise from

the testing side. These issues can be industry driven changes,

changes to the type of testing required, the introduction and

availability of new methods or consumables, feedback from users

of the Standard or personal experience when using the method

in the laboratory. The Standards are reviewed and it is then the

decisionmust bemadeonwhat todowith theStandard. They canbe

reconfirmed (no changes), withdrawn or revised (the Committee

evaluates and revises the Standard).

When a Standard is released for public comment it is crucial that

the users of the Standard read it and make any relevant comments.

All of these comments are documented and must be discussed by

the committee at a meeting. The discussion outcome is also docu-

mented and any changes required are made and eventually the

Standard is printed. It is not an easy process. Writing an unambig-

uousStandard takes timeanda lot of discussion. Thinkback to those

tasks you had at TAFE or university when you had to write instruc-

tions on how to cook a piece of toast. It is often the little things that

can be overlooked, the things you may take for granted. There are

certain guidelines that have to be followed and are set out by

Standards Australia. The method steps are there to follow so that

everyone using the Standard is doing the same thing therefore

creating a uniform procedure that should give the same results no

matter where you are. Standard procedures are important and it is

the basis of the quality systems in laboratories.

So please comment if you feel there is something you need to bring

up when a Standard goes out for Public Comment. Even if it just a

typographical error or something you think is wrong-COMMENT.

Your company can subscribe to StandardsWatch so you are alerted

when a Standard relevant to your area is released or open for public

comment.

Thank you to the ASM and my employer (ALS) for supporting me

to be a committee member on FT020 Water Microbiology. It is

appreciated and it is only through organisations like the ASM

and generous employers that Standards Australia committees can

operate. All committee members are volunteers and must have

the relevant qualifications and knowledge to ensure that these

Standards can continue to be developed and maintained.

2016 ASM Communication Ambassador ProgramThe ASM is looking for teams of student and ECR members to

volunteer as part of a roster of Communication Ambassadors within

the society, who will:

(1) Be in charge of ASM’s online communication channels togetherwith a team of ASMmembers, bolstering the society’s networksby sharing your story with scientists, industry, media membersand policy makers;

(2) Effectively represent the society by using these channels toshowcase yourwork, reveal personal insights intoyour scientificcareer, and serve as a public advocate for microbiology.

We are looking for enthusiastic members who are keen to contrib-

ute. To apply for the 2016 program, please submit an Expression of

Interest (maximum 200 words) to [email protected] by 31

March 2016 and highlight the following:

* Your name, institution, qualifications, area of interest withinmicrobiology, and ASM membership status;

* Your strategy for broadening ASM’s online presence and collab-orating with other communication ambassadors;

* What conferences, scientific meetings, or events you can providesocial media coverage for in 2016.

Successful applicants for the 2016 Communication Ambassador

program will receive a $100 discount for registration to the 2016

ASM Annual Scientific Meeting (3–6 July 2016, Perth Convention &

Exhibition Centre).

ASMAffairs


Stopping dengue: recent advances and newchallengesGayathri Manokaran, Kirsty McPherson and Cameron P Simmons

Symposium 16 November 2015

Peter Doherty Institute for Infection and Immunity,

Melbourne

Dengue remains a major problem throughout the world with an

estimated 30% of the world’s population at risk of infection. In the

past few years, major advances have been made across virology,

clinical insights, vaccines and mosquito control strategies. The first

of its kind to be ever held in Australia, the International Dengue

Symposium 2015 showcased some of the best science and clinical/

public health research being undertaken in the field.

Proudly organised by The Peter Doherty Institute for Infection and

Immunity, under the leadership of Professor Cameron Simmons,

this symposium brought together 130 national and international

dengue researchers. The organising committee acknowledges the

support of ASM as an important partner to this meeting, together

with generous support from The University of Melbourne, The

Royal Melbourne Hospital, the Oxford University Clinical Research

Unit, Vietnam and the DUKE-NUS Graduate Medical School,

Singapore. Industry sponsors included Sanofi Pasteur, BioTools,

VWR International, In Vitro Technologies, Pacific Lab Products,

Interpath Services and Alere Global.

The formal symposium was opened by Professor Cameron

Simmons. This was followed by a series of 21 brief talks spanning

across four main themes of dengue research: Virology, Public

Health, Clinical Research and Immunology/Pathogenesis. Short

breaks for networking were scheduled in between each of these

four sessions.

All of the invited speakers had considerable international profiles in

dengue and hence the quality of the presentations was very high.

Associate Professor Sheemei Lok discussed the structural basis

behind how a dengue-2-specific human monoclonal antibody

effectively protects mice from dengue infection due to its ability

to ‘lock’ the dengue envelope proteins while blocking the binding

of enhancing antibodies. Following that, Professor Ooi Eng Eong

described the novel finding of a viral RNA-host protein interaction

that allows dengue virus to evade the host immune response- this

work was recently published in the prestigious journal, Science.

Ahighlight of this symposiumwas the ‘PublicHealth’ session,which

was the first of its kind in Australia, with speakers from various

companies includingSanofiPasteur andTakeda, coming together to

discuss their respective dengue vaccines, thereby providing atten-

dees with a useful summary of all the dengue vaccines currently in

clinical development.

Novel and highly innovative medical technologies were next dis-

cussed in the ‘Clinical Research’ session, with Dr Jenny Low

explaining how in vivo imaging technologies such as positron

emission tomography are being optimised presently to track den-

gue virus progression in a mouse model. Dr Sophie Yacoub gave a

summary on changes in microvasculature of dengue patients upon

infection and how these disturbances might precede the develop-

ment of severe disease.

In the next session, Dr Laura Rivino discussed their work on

elucidating the human cellular responses during dengue infection

and how both CD4+ and CD8+ T-cell subsets remain enriched in

the skin of dengue patients up to 5months post infection. Professor

Cameron Simmons then described an interesting dengue neutra-

lisation assay developed in Vietnam using patients’ viraemic blood

and infected mosquitoes as a readout of the efficacy of monoclonal

antibodies to efficiently neutralise dengue virus. Next, Dr David

Muller introduced the concept of nanopatches that allow vaccine

delivery in a needle-free manner.

In closing, the sponsors were wholeheartedly acknowledged for

their contributions by Professor Cameron Simmons and their

support was instrumental in allowing this symposium to be a

success. Not only was the stellar line-up of talks highly educational

but importantly, this symposium provided a networking opportu-

nity for everyone interested in meeting the challenge of reducing

dengue burden throughout the world.

ASMAffairs


Confirmed Plenary speakers

Annual Scientific Meeting and Trade Exhibition

www.theasm.org.au www.westernaustralia.theasm.org.au

Professor Peter Hawkey University of Birmingham Nosocomial infection control and antibiotic resistance

Professor Dan Andersson Upsalla University Environmental pollution by antibiotics and its role in the evolution of resistance

Assoc Prof Susan Lynch University of California San Francisco Colitis, Crohn's Disease and Microbiome Research

Professor Anna Durbin Johns Hopkins Dengue and vaccines

Dr Brian Conlon Northeastern University, Boston Drug discovery in soil bacteria

Watch this space for more details on the scientific and social program, speakers, ASM Public Lecture, workshops, ASM awards, student events, travel awards, abstract deadlines and much more..

Perth, WA A vibrant and beautiful city located on the banks of the

majestic Swan river. Come stay with us in WA and experience our world class wineries and restaurants, stunning national parks, beaches and much more..

NOW CONFIRMED! 2016 Rubbo Oration Professor Anne Kelso CEO NHMRC

As with previous years, ASM 2016 will be co-run with EduCon 2016: Microbiology Educators’ Conference

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