DIAGNOSIS OF GYRODACTYLUS (MONOGENEA; FISH IN … · Other Gyrodactylus species commonly found on salmonids in Europe include Gyrodactylus derjavini Mikailov, 1975, Gyrodactylus truttae

FRS Report No 07/02

Not to be quoted without prior reference to the authors

Fisheries Research Services Report No 07/02 DIAGNOSIS OF GYRODACTYLUS (MONOGENEA; PLATYHELMINTHES) INFECTING SALMONID FISH IN NORTHERN EUROPE

C M Collins, T A Mo, K Buchmann and C O Cunningham

March 2002 Fisheries Research Services Marine Laboratory Victoria Road Aberdeen AB11 9DB

Contents Introduction ................................................................................................................. . 1 Mode of pathogenicity ....................................................................................... . 1 Spread of parasite ............................................................................................. . 2 Treatment and Prevention ................................................................................ . 2 Legislative Control ............................................................................................ . 2 Identification ...................................................................................................... . 2 Diagnostic Procedures ................................................................................................ . 3 Sampling ..................................................................................................................... . 3 When to sample ................................................................................................ . 3 Collection of data .............................................................................................. . 3 Catching fish for examination ........................................................................... . 3 Numbers of fish to be examined ....................................................................... . 3 Condition of fish ................................................................................................ . 4 Sampling Gyrodactylus from live fish ............................................................... . 4 Sampling Gyrodactylus from fixed/preserved fish ............................................ . 4 Examination of Fish .................................................................................................... . 5 Diagnosis Using Morphological Means ...................................................................... . 9 Slide preparation ............................................................................................... . 11 Analysis of morphological characters ............................................................... . 14 Anchors ................................................................................................. . 20 Ventral bar ............................................................................................. . 22 Dorsal bar .............................................................................................. . 22 Marginal hooks ...................................................................................... . 23 Cirrus ..................................................................................................... . 24 Host ....................................................................................................... . 24 Morphological Analysis of Gyrodactylus species Commonly Found on Salmonids In Europe ................................................................................................................ . 24 Gyrodactylus "Quick" Identification Checklist ............................................................. . 34 Diagnosis Using Molecular Means ............................................................................. . 36 Preparation of samples ..................................................................................... . 37 The V4 region of the small subunit ribosomal RNA gene ................................ . 37 PCR amplification of the V4 region ...................................................... . 37 Visualisation of PCR products on gel ................................................... . 37 Restriction digest of the V4 region ....................................................... . 38 Generation of probes ............................................................................ . 38 Preparation of membranes for hybridisation ........................................ . 41 Hybridisation of labelled probes to membrane-bound samples ........... . 42 Detection of labelled probes bound to membrane ............................... . 42 Analysis ................................................................................................. . 42 Ribosomal RNA internal transcribed spacer (ITS) ........................................... . 44 PCR amplification of ITS ....................................................................... . 47 Visualisation of ITS PCR products on gel ............................................ . 47 Restriction enzyme digest of the ITS .................................................... . 47 Visualisation of restriction digest products on gel ................................ . 48 Amplification of the ITS1 and ITS2 regions of Gyrodactylus Ribosomal DNA .................................................................................... . 52

Amplification of ITS1 ............................................................................. . 52 Amplification of ITS2 ............................................................................. . 52 Restriction enzyme digest of ITS1 ........................................................ . 52 Restriction enzyme digest of ITS2 ........................................................ . 53 Sequence analysis of the V4 and ITS ribosomal DNA regions ....................... . 54 Integration of Morphological and Molecular Diagnostic Methods .............................. . 56 Outlined method for integrated identification of Gyrodactylus parasites ......... . 57 Case Studies ............................................................................................................... . 59 G. salaris/G. teuchis ......................................................................................... . 59 G. salaris variant ............................................................................................... . 59 G. salaris/G.thymalli .......................................................................................... . 60 Additional Methods for the Identification of Gyrodactylus species ............................ . 60 Release of opisthaptoral attachment structures from tissue ............................ . 60 Scanning Electron Microscopy (SEM) .............................................................. . 63 Statistical classifiers .......................................................................................... . 64 Chaetotaxy ........................................................................................................ . 64 References ................................................................................................................ . 65

Diagnosis of Gyrodactylus

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DIAGNOSIS OF GYRODACTYLUS (MONOGENEA; PLATYHELMINTHES) INFECTING SALMONID FISH IN NORTHERN EUROPE

C M Collins1, T A Mo2, K Buchmann3 and C O Cunningham1

1 FRS Marine Laboratory, PO Box 101, Victoria Road, Aberdeen, AB11 9DB 2Fish Health Section, National Veterinary Institute, PO Box 8156 Dep, 0033 Oslo, Norway 3Department of Fish Diseases, Royal Veterinary and Agricultural University, Bülowsvej 13,

Frederiksberg C, Denmark

INTRODUCTION Gyrodactylus parasites are one of a number of parasites found infecting freshwater salmonids in Europe. They are small (300-800 µm), ectoparasites, with an attachment organ, the opisthaptor, at one end and mouthparts at the other (see Williams and Jones, 1994). They are remarkable in that they give birth to live young, which already have a developing embryo, in a ‘russian doll’ arrangement. For the most part they are thought to be relatively benign and are not a cause for concern with respect to fish health and mortalities. The exception to this is Gyrodactylus salaris Malmberg, 1957. G. salaris has only been found in Europe (Malmberg, 1993). It occurs in the Baltic regions and seems to coexist with Baltic strains of salmon (Salmo salar) without problems. However, the parasite is extremely pathogenic to many Atlantic strains of salmon. Since its introduction to Norway in the 1970’s it has resulted in the decimation of salmon stocks in numerous Norwegian rivers (Johnsen and Jensen, 1991, Johnsen et al., 1999). The Scottish salmon stock so far tested was shown to be equally susceptible (Bakke and Mackenzie, 1993). G. salaris can survive and reproduce on several different salmonids, such as rainbow trout (Oncorhynchus mykiss), Arctic char (Salvelinus alpinus), North American brook trout (Salvalinus fontinalis), grayling (Thymallus thymallus), North American lake trout (Salvalinus namaycush) and brown trout (Salmo trutta) (in declining order of susceptibility) (Bakke et al., 1991a; 1991b; 1991c; 1992a; 1992b; 1996; 1999). Other Gyrodactylus species commonly found on salmonids in Europe include Gyrodactylus derjavini Mikailov, 1975, Gyrodactylus truttae Gläser, 1974, Gyrodactylus teuchis Lautraite, Blanc, Thiery, Daniel and Vigneulle, 1999, and Gyrodactylus thymalli Zitnan, 1960. These species show varying preferences for different host species (Buchmann and Uldal, 1997). The possibility of finding several different species of gyrodactylids on the same host species, combined with the particular pathogenicity of G. salaris to Atlantic salmon, makes correct diagnosis of Gyrodactylus parasites infecting freshwater salmonids essential. Mode of Pathogenicity During the course of feeding and attachment to the fish, the Gyrodactylus parasite damages the host epidermis. Holes in the epidermis are caused by the hooks and anchors of the attachment organ, and ulcers are generated by enzymatic digestion (Mo, 1994). Infection with thousands of Gyrodactylus parasites, as is seen in the case of Atlantic salmon parr infected with G. salaris, results in severe damage to the fish epidermis and loss of ability to osmoregulate properly. This is thought to be the principal cause of host mortality. Secondary infections of the epidermal lesions with bacteria or fungi may also play a significant role in the pathogenicity of Gyrodactylus.


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Spread of Parasite Gyrodactylus salaris has spread between rivers and farms mainly through the transport and restocking of live fish (Mo, 1994; OIE, 2000). Even though G. salaris is a freshwater parasite and cannot survive full-strength sea water, it has been shown to survive for up to 240 and 42 hours at 10‰ and 20‰ salinity, respectively, and as such there is the possibility of spread of the parasite by fish migration between adjacent rivers via low salinity fjords (Soleng and Bakke, 1997; Soleng et al., 1998). The use of fishing tackle which has not been properly dried or disinfected is also a theoretical source of parasite transmission, but of much lower risk than fish movements. Treatment and Prevention Treatment of Gyrodactylus infections in fish hatcheries and onshore farms can be undertaken using a variety of chemical or drug regimes (Buchmann, 1997; Crigel et al., 1995; Lindenstrøm and Buchmann, 1999; Santamarina et al., 1991; Schmahl, 1993; Schmahl and Taraschewski, 1987; Soleng and Bakke, 1997; Soleng et al., 1999; Tojo and Santamarina, 1998 ; Tojo et al., 1992; 1993). In Norway, salmonid hatcheries were completely cleared and dried for a period to guarantee that all G. salaris had been removed. In natural watercourses, removal of Gyrodactylus parasites is more difficult. In Norway, the drastic action of rotenone treatment, killing off all fish hosts, has been undertaken to eradicate G. salaris in some rivers (Johnsen and Jensen, 1991; Johnsen et al., 1999; Mo, 1994). This is environmentally harsh and can only be carried out in selected rivers that are short with few tributaries, and with a low level of species diversity. Rotenone treatment in other European countries may not be feasible due to the geographical or biological nature of their river systems. Legislative Control Within the European Community, G. salaris is placed on list III of Directive 91/67/EEC. This list contains diseases that have a significant economic impact in certain circumstances and may warrant national control measures. Under Commission Decision 96/490/EC on certain protective measures with regard to Gyrodactylus salaris in salmonids, susceptible species cannot be moved to areas of the EC that have been shown to be free of G. salaris unless the zone of origin has undergone a period of testing and has also demonstrated freedom from G. salaris. Monitoring for the parasite and strict control of the movement of stocks between rivers has been an integral part of the Norwegian strategy to prevent the spread of G. salaris. Monitoring programmes to demonstrate freedom from G. salaris are being planned or executed by an increasing number of countries. These programmes can be complicated by difficulties in identifying Gyrodactylus specimens to species level. Identification Gyrodactylus species are usually identified using morphological characteristics, principally those of the attachment organs, but also in more detailed studies, the protonephridia (Malmberg, 1970). Some species are easily and quickly differentiated from each other. However, others are very similar morphologically and differentiation requires detailed analysis of the characters used in identification. These characters can also vary depending on environment, host species, and water temperature (Mo, 1991a; 1991b; 1991c). As a result, a high level of experience is required for accurate identification of Gyrodactylus species. In addition to the variability in morphology, problems for identification can be encountered during preparation of the parasites for microscope examination. Preparation of


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slides can be time consuming and difficult, the characters used in identification needing to be orientated properly for reliable diagnosis. This is especially difficult in the case of preserved specimens, a situation that is usually necessitated during the collection of samples for diagnostic laboratories. Developments in molecular techniques have resulted in their application to parasite diagnostics. Molecular criteria can be more objective than morphological criteria for species identification, and can easily be performed by personnel after a minimum of training. The general methodology set out in this guide can also be applied to Gyrodactylus parasites infecting other fish families, but details of measurements and nucleotide sequences are given only for species found to date on aquacultured salmonids in Europe.

DIAGNOSTIC PROCEDURES Sampling When to sample Outbreaks of gyrodactylosis, caused by G. salaris can occur at any time, but are most common in spring and in periods when the water temperature is 7-17ºC (OIE, 2000). Collection of data Information regarding the sampling date, water temperature, host species, size/age of host, sampling locality, source of fish if restocked, and water chemistry, should be recorded if available. Minimum data of host species and sampling location should be recorded. Suitable labels should be given to fish and/or tubes so that different samples can be unambiguously identified. Catching fish for examination Electro-fishing, bow net, scap net, small trawl and seine are all acceptable methods for catching fish for Gyrodactylus examination. Methods that do not cause damage to the external surface of the fish, such as electrofishing and sport fishing, are preferable. Electrofishing is preferable when sampling fish in the wild. Note: Nets may damage the fish and cause Gyrodactylus specimens to drop off (Malmberg, 1970). Numbers of fish to be examined A minimum of 30 fish per site should be sampled and examined. This will give a detection rate of 95% when the prevalence of Gyrodactylus is 10%. The whole fish must be examined to give 100% sensitivity. If only the fins are examined the sensitivity is reduced and consequently the number of fish to be examined must be increased. A Gyrodactylus species may occur in much lower prevalence than 10%, for example on resistant host strains or low susceptible host species. In these cases the number of sampled fish should be significantly higher than 30.


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Condition of fish Fish must be live when collected. Gyrodactylus parasites often leave a host soon after it dies. Dead fish, transported on ice, are not acceptable for Gyrodactylus examination. The parasites soon die if not covered in water, and disintegrate quickly. In such cases, if no parasites are found, either on the fish or after dropping to the bottom of the container, it cannot be concluded that the fish were uninfected (OIE, 2000). Sampling Gyrodactylus from live fish (see Malmberg, 1970; OIE, 2000) • Avoid handling the fish as much as possible as this may displace parasites from the

surface of the host, ie, use a net. • Gyrodactylus specimens may die or detach from the host if water quality or chemistry

is changed. Therefore maintain the fish in water from the site at which they were caught.

• Fill an additional container containing site water only, for subsequent use during fish examination.

• The density of fish in the container should not exceed one 15 cm fish per litre of water. This is to prevent too great a change in water quality due to the build up of excretory (and other) products from the fish. If not examining fish on site, the volume of water allowed per fish may also be influenced by transportation time.

• Different fish species should be maintained separately as mixing host species may initiate detachment of the parasites. This may be induced by the presence of fish excretory products in a small quantity of water.

• Keep the water containing the fish cold (eg by use of shade, cold storage room or refrigeration). Do not add ice or water.

Sampling of Gyrodactylus from fixed/preserved fish (see OIE, 2000) Fish can be fixed in formaldehyde or preserved in alcohol. • The concentration of formaldehyde after fixation should not be lower than 4% (v/v)

(10% (v/v) formalin). The formaldehyde concentration should be 8-10% (v/v) (20-25% (v/v) formalin) before adding the fish, because water is freed from the fish during fixation.

For detection of Gyrodactylus specimens on the fish, formaldehyde fixed fish are preferable to ethanol preserved, because with the former, the parasites become opaque and easier to detect. However, formalin fixed parasites cannot be readily used subsequently for molecular diagnostic techniques. Therefore, if molecular diagnostic techniques are to be used, ethanol preservation is preferred. Ethanol is also less hazardous to workers than formalin fixatives. • The concentration of ethanol after preservation should not be lower than 70% (v/v).

Again, water, freed from the fish following preservation must be allowed for. If the ethanol concentration is lower than 70% (v/v), the mucus and epidermis may disintegrate and Gyrodactylus specimens, even if preserved, may drop off. Conversely, the ethanol concentration should not be too high, as the fish tend to shrink, making them more difficult to examine. A concentration of 80-85% (v/v) ethanol before addition of the fish is recommended.


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• Ethanol preserved fish should preferably be stored in a cold room, especially if long time periods elapse between collection and examination.

• The fish should be fixed or preserved in relatively large bottles that provide excess space and fixative/preservative (5:1 ratio of volume of fixative:volume of sample).

• The opening of the bottles should be wide to avoid the possibility of scraping off Gyrodactylus specimens when the fish are taken out for examination.

• Containers and fish should be stored in a horizontal position during fixation/preservation. This reduces the amount of distortion in the fish and makes later examination easier. When fish have been fixed/preserved for 2-3 days, the bottles can be stored in a vertical position.

Examination of fish (see Malmberg, 1970, OIE, 2000) Infection of Atlantic salmon with G. salaris usually results in gyrodactylosis, which is detrimental to the health of the fish, and can result in mortality. In cases of gyrodactylosis, the fins, especially the dorsal and pectoral fins, are most commonly infected but parasites can occur on all epidermal surfaces, including the body, nostrils, gills and mouth cavity (OIE, 2000). In these cases, scrapings can be taken from the surface of the fish and these will usually contain specimens of the parasite. However, occasionally in the case of G. salaris infecting Atlantic salmon, and often in the case of other Gyrodactylus species infecting salmonids, the numbers of parasites present on a fish can be very low, with single parasite infestations common. In these situations, the chances of detecting parasites in scrapings are limited, and examination of the whole fish, as outlined below, is necessary (OIE, 2000). In the case of monitoring programmes, it is recommended that the whole fish be examined. This can be difficult and very time consuming for fish larger than about 20 cm in length and in these cases only the fins are examined. All fins from a fish should be examined. It is important to note that this will reduce the detection of parasites from fish with low infection levels. In addition, it has been shown that certain species of Gyrodactylus prefer different sites on the fish, eg G. thymalli seems to prefer the body surface rather than fins, and as such, sampling fins only will decrease detection of this parasite (Sterud et al., 2002). Changes in site preference of the parasite can also change over time in a given host/parasite system, possibly in relation to host response. • Fish should be examined individually under a binocular dissecting microscope with

good illumination. The fish can be examined under 12-20x magnification. A light source which can be directed in the desired manner at the specimen is required. When investigating light-refracting parts of the fish, eg parts of the skin and pharynx, an obliquely directed beam of light will create a better contrasting effect.

• The fish should be transferred, using a net, to a suitable sized box, filled with water from the sampling site. If the dissecting microscope is illuminated from above, the bottom of the box should be black. This will increase the contrast and the parasites will be more easily detected.

• Live parasites are more easily detected by their movements, so disturbing light refraction on the skin of the fish should be avoided.

• The fins of small, unfixed fish, less than 10cm, can be studied using illumination through the bottom of the box. Gyrodactylus specimens on the fins can easily be observed in this way.

• Catch the fish by means of a claw forceps. Place the forceps just behind the head. Avoid touching the fish with hands.

• Holding the fish by means of the forceps, insert a preparation needle into the brain through the upper part of the eye. This will kill the fish instantly. In addition, this


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method has the advantage that it avoids blood shed into the water, which might affect the parasites.

• The whole surface of the fish, including the gills and mouth cavity, must be examined. It is best to use two sets of forceps for this process.

• Each fish should be examined for at least five minutes. However, this depends on the size of the fish; larger fish will take longer to examine.

As an alternative to examining the fish as a whole, it can be dissected into different parts prior to examination (Malmberg, 1970): • Holding the fish with forceps, cut off the fins using a curved pair of scissors and place

them directly in a separate dish with site water. • Still holding the fish with the forceps, insert the pointed blade of a pair of scissors into

the fish mouth and sagitally divide the skull and the lower jaw. • Decapitate the fish, cutting just behind the opercula, directly placing the two halves of

the head in a separate dish, covered with site water. • Place the body of the fish in a separate dish with site water. • Cut off the gill arches from the halves of the head, one by one, and transfer them to a

separate dish with site water. • Place the above dishes in cool place (4ºC) while awaiting examination. As an alternative to killing the fish, the fish can be anaesthetised with MS222 (3-aminobenzoic acid ethyl ester), chlorobutanol, benzocaine, or other suitable anaesthetics. The fish is then examined for the presence of parasites under a dissecting microscope. Small, anaesthetised fish can be placed under the microscope in a suitably sized container containing anaesthetic. Examination of the fish should be carried out as quickly as possible to ensure that fish subsequently recover from the anaesthetic. In general, Gyrodactylus parasites are not affected by the anaesthetic and only a very low percentage will detach from the fish. The liquid and the base of the container can be examined for parasites after the fish has been removed. Fixed or preserved fish should be studied in a similar way under a dissection microscope with illumination from above. Gyrodactylus specimens turn almost white when fixed in formaldehyde, while ethanol preserved specimens are slightly opaque (OIE, 2000) (see Fig. 1). • Before examination, fish that have been fixed in formaldehyde solution should be

rinsed in tap water. This can be done by placing the fish in a new container under a tap and letting the water flow gently (so as not to detach parasites from the fish) through the container for a couple of hours. The remaining fixative, including the bottom sediment in the original transport container, should be examined separately for the presence of Gyrodactylus specimens that may have washed off during fixation and transportation. Ideally, for safety purposes, the dissecting microscope should be placed on a suction bench with downwards outlet to avoid inhalation of evaporated fixative.

Figure 1 (a-d) shows both fresh and ethanol preserved specimens of Gyrodactylus parasites, attached to fish fins.


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Figure 1b. Ethanol preserved fish fin with attached Gyrodactylus parasite (see arrow)(x125)

Figure 1a. Fish fin with live Gyrodactylus parasites attached (see arrow)(x125)


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Figure 1c. Ethanol preserved fish fin with attached Gyrodactylus parasite (see arrow)(x170)

Figure 1d. Ethanol preserved fish fin with attached Gyrodactylus parasite (see arrow)(x220)


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Diagnosis using morphological means (see Malmberg, 1970, OIE, 2000) Gyrodactylus specimens are identified individually. Several Gyrodactylus species may occur on a single fish. It is therefore recommended that several Gyrodactylus specimens be prepared for identification, preferably from different sites on the fish. A minimum of five parasites should be examined per fish, but these parasite preparations must be of suitable quality to allow identification by morphological means. Therefore specimens should continue to be examined until five specimens have been identified (or are of sufficient quality to be identified later) by morphological means, or until all parasites on fish have been examined, whichever comes first. The classical method of classifying and identifying species is based on examination of morphological characters of taxonomical importance. The morphological characters used in Gyrodactylus species descriptions are shown in Figure 2. Preparation of Gyrodactylus parasites for morphological examination is described below. Malmberg’s ammonium picrate-glycerine (APG) method for preparing whole mounts of small opisthaptor worms (Monogenea) is superior to other methods (OIE, 2000). The APG is prepared by mixing (1:1) one part saturated ammonium picrate solution and one part glycerine/glycerol (puriss). Ammonium picrate is difficult to dissolve. Unless ammonium picrate crystals remain undissolved at the bottom of the mixing bottle, the solution will not be saturated solution. To ensure saturation, shake the bottle regularly during one week prior to use (Malmberg, 1957). Warning: when dry, ammonium picrate is explosive.


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Figure 2. Morphological features used in the identification of Gyrodactylus parasites are arrowed. The opisthaptoral hardparts, shown in greater detail in figure 1b, are used to differentiate between Gyrodactylus parasites at the species level.


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The specimen can be divided into two if required; the opisthaptor can be retained for further morphological examination, and the body can be used in molecular diagnostic techniques. The opisthaptor may be prepared as described for whole mounts. This has considerable advantages, as both morphological and molecular analysis can be carried out on the same specimen. However, it has the disadvantage of being a technically demanding method, with a high risk of losing at least one part of the parasite during dissection and transfer to slides or tubes. Slide preparation (see Fig. 3) • Place a drop of water on a microscope slide (76 x 26mm). If the parasite is fixed,

then it is not necessary to place a drop of water on the slide, the parasite can be placed directly on the slide in its fixative.

• Transfer a single parasite to the water drop using needle or fine forceps. • Place a coverslip gently on top, lowering the coverslip onto the slide at an angle to

avoid air bubbles. • At this stage, examine the specimen to see if it is in the correct orientation. If not, the

coverslip can be removed and the parasite reoriented with the tip of forceps or needle. Sometimes moving the coverslip gently from side to side with the tip of forceps, while observing the specimen under a dissecting microscope, can improve the orientation of the parasite. However, this can also easily result in damage to the parasite.

• Absorb excess water from under the coverslip by placing a piece of filter paper at the edge of the coverslip. This causes the worm to be compressed on to one plane so that the morphological features of the attachment organ or opisthaptor can be observed. This method avoids excessive force, which might damage the parasite, for example by pushing down the coverslip from above with a forceps or needle. Alternatively, if preparing several slides for examination, each parasite can be left at this stage until all have been placed under coverslips. Then, starting with that prepared first, and continuing sequentially, the APG can be added. Usually, by the time APG is added to each slide, the excess water will have evaporated, leaving the worm compressed.

• Add a small drop of APG to the edge of the coverslip. The parasite will be fixed as the yellow APG-solution penetrates the space between the slide and the coverslip.

• Absorb excess APG from under the coverslip by placing a piece of filter paper at the opposite edge of the coverslip.

• Label the slide with a unique identifier for the parasite. • Permanent attachment of the coverslip, if required, can be obtained by adding a

small drop of nail polish or similar substance to each corner of the coverslip. • Examine the opisthaptor of the parasite. The marginal hooks, anchors, ventral and

dorsal bars, are also often visible in well developed embryos and can sometimes appear clearer than those in the adult.

Preparation of live parasites will give the best preparations. Fixed or preserved parasites are prepared for morphological identification as above. However, they are harder to depress onto one plane and identification may prove difficult (OIE, 2000). Ethanol preserved parasites can be placed in water for 1-2 hours or longer before preparation. The parasites will rehydrate and become easier to compress. If using a high power objective with immersion oil, the parasite should be placed as close to the centre of the coverslip as possible. This is to prevent immersion oil seeping between the coverslip and slide and destroying the preparation (Malmberg, 1970).


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If examining the excretory system of Gyrodactylus parasites, a phase contrast microscope must be used. It is not possible to see the excretory system in clear detail under bright field microscopy. Other soft body parts such as the pharynx, ovary and muscles can only be studied under phase contrast. Phase contrast microscopy is also superior to bright field microscopy for study of the opisthaptoral hard parts (Malmberg, 1970). Formalin fixed specimens, stained in eosin, Gomori's trichrome stain, Ehrlich or Mayers stain, among others, and mounted in Canada balsam, glycerine-gelatine or Depex, can be examined under bright field microscopy. These methods provide good visualisation of the ventral bar, and the ventral bar membrane (Buchmann, unpublished observations).


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Preparation of Gyrodactylus parasite for morphological examination

Add Ammonium Picrate Glycerine (APG)

Place parasite in drop of water onslide. If parasite is preserved, it is not

necessary to place in drop of water.

Remove excess liquid withfilter paper

Add coverslip

Fix coverslip to slide

Remove excess APG withfilter paper

Preparation of Gyrodactylus parasite for morphological examination





Add coverslip















Add coverslip



Add coverslip

Fix coverslip to slideFix coverslip to slide



Figure 3. Slide preparation of Gyrodactylus parasites, mounted in ammonium picrate-glycerine, for morphological examination of hard parts.


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Analysis of morphological characters When analysing morphological characters, two approaches can be taken. The overall shape of different characters is noted, and the measurement of certain characters is recorded. The pharynx, excretory systems, cirrus, anchors, dorsal and ventral bars and marginal hooks of Gyrodactylus parasites are of taxonomic value (Fig. 2) (Malmberg, 1970). The protonephridial system can be used to divide Gyrodactylus species into subgenera. Keys for dividing Gyrodactylus species into subgenera, based on the morphology of the protonephridial system, were described by Malmberg (1970). Species within a subgenus can be further divided into smaller species groups based on the size and shape of the anchors and ventral bar. Species discrimination in Gyrodactylus is mainly based on the shape of the marginal hooks of the opisthaptor, but all opisthaptoral structures are taxonomically important (Malmberg, 1970; 1993). It is practically impossible to separate very similar species by means of character measurements (Malmberg, 1970). Hard parts of Gyrodactylus parasites are more useful in distinguishing between different taxonomic groups than the soft parts (Malmberg, 1970). The features of the individual opisthaptoral characters used in morphological analysis (anchors, marginal hooks, ventral bar and membrane and dorsal bar) can be subdivided into different regions for descriptive purposes and to make comparisons between different species easier. Figure 4 represents a common subdivision of Gyrodactylus opisthaptor characters. Measurements can be taken directly from a slide under the microscope using a micrometer eyepiece, or from a projected image (Mo, 1993). The characters and measurement criteria described below have been taken from Malmberg 1970. Points from which measurements are taken are shown in Figure 5a. These measurement criteria are now adopted by many authors when describing Gyrodactylus species, and so help to standardise the descriptions.


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Figure 4. Diagrammatic illustration of the subdivisions of the opisthaptoral hardparts of Gyrodactylus, used in species discrimination. This figure has been adapted from Shinn et al., 1995. (A) Anchor; a.p.=point, a.s.= shaft, a.r.=root a.j.=dorsal bar attachment point; a.k.=ventral bar attachment point; a.l.=indentation marking lower edge of the ventral bar attachment point (B) Marginal hook: sickle proper: h.p.=point; h.a.=shaft of the sickle proper; h.t.=toe; h.h.=heel; h.t. +h.h.=foot/base of the sickle proper; h.s.=shaft; h.f.=indentation noted in toe of certain species; h.g.=aperture. (C) Ventral bar: v.p.=processes; v.d.=transverse depression; v.o.=median portion; v.m.=membrane; v.r.=medial ridge.

a.s

a.r

a.ja.ka.l

a.p

h.p

h.a

h.th.h

h.s

h.fh.g v.dv.o

v.m

v.r

hood-like

anchor

filament

marginal hook

Indentation

v.psickle

membran


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(a)

l.si.f

Figure 5a. Diagrammatic illustration of measurements taken on anchors and marginal hooks. Abbreviations: (A) Anchor: l.a.= total length of anchor; l.a.r. = length of anchor root; l.a.s. = length of anchor shaft; l.a.p. = length of anchor point (B) Marginal hook: l.m.h. = total length of marginal hook; l.si. = length of sickle; l.h. = length of handle; w.d.s. = width (distal) of sickle; w.p.s. = width (proximal) of sickle; l.si.f. = length of sickle filament loop. Figure A adapted from Mo (1993); figure B adapted from Shinn et al. (1995); Malmberg (1970).

l.a.s

l.a.r

l.a

l.a.p

A

l.m.h

l.h

l.s.i

w.p.s

w.d.s

B


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Figure 5c. Features of shape, relative size, and orientation of certain characters of the marginal hooks, used in the morphological identification of Gyrodactylus parasites, are illustrated in figure 5c. Drawings of marginal hooks are taken from;1 Ergens (1983), 2 Cunningham et al, (2001). G. thy: Gyrodactylus thymalli, G.s: G. salaris, G. teu: G. teuchis, G.d: G. derjavini, G.tru: G. truttae.

G. d G. tru

G. s G. teu G. thy

(c) 1 1

1

2

1

(b)

G.s G. d G. tru

Figure 5b. The orientation of the anchor root in relation to the anchor shaft, and the curvature of the anchor point in relation to the anchor shaft, can be helpful in identification of Gyrodactylus species. Drawing of anchors taken from 1 Ergens (1983).

1 1 1


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(d)

Figure 5d. (a) Length of ventral bar longer than maximal distance between processes of ventralbar; (b) Maximal distance between processes of ventral bar longer than length of ventral bar.l.v.b. = length of ventral bar; m.d.p.v.b. = maximal distance between processes of ventral bar;t.w.v.b. = total width of ventral bar; l.p.v.b. = length of processes of ventral bar; b.w.v.b. = basalwidth of ventral bar; m.w.v.b. = median width of ventral bar; l.v.b.m. = length of ventral barmembrane. Figure taken from Malmberg (1970).

Figure 5e. The size and shape of the ventral bar processes are important in Gyrodactylus species identification. The perceived angle of the processes, in relation to the median portion of the ventralbar (marked in red), is sometimes used in species identification, but in general it is not considered very useful. * G. derjavini and G. truttae can display either type of angle shown here in red. Drawings of ventral bars and membranes were taken from 1 Ergens (1983); 2 Cunningham et al. (2001);3 Mo (1993). G. thy: Gyrodactylus thymalli, G.s: G. salaris, G. teu: G. teuchis, G.d: G. derjavini, G.tru: G. truttae.

(e)

G.s G.teuG.thy G. truG.d

2 22 3 1

* *


19

Figure 5f. Different types of dorsal bars. l.d.b. = total length of dorsal bar; m.w.d.b. =median width of dorsal bar. Figure taken from Malmberg (1970)

(f)


20

It must be remembered when comparing measurements within and between Gyrodactylus species, that different methods of specimen preparation, different types of microscopy and different methods of measuring can cause differences in results. This is especially true in the case of the anchor roots, ventral bars and marginal hooks (Malmberg, 1970). Measurements can also vary depending on water temperature (they tend to be larger during the colder part of the year), host species, and host size. For this reason, and the occurrence of species which have very few and very slight differences between them, an adequate number of measurements must be taken. Note: While each of the opisthaptoral parts described below can be useful in Gyrodactylus species identification, it is the overall picture given by all the parts which is important in diagnosis. Caution should be used if comparing individual parts between different species. Anchors Note: Anchors are also referred to as hamuli (singular hamulus) by many authors. The term “anchor” will be used in this Guide. Anchor shape can be used to distinguish between many, but not all, Gyrodactylus species. The anchor can be divided into three main parts; root, shaft and point (Fig. 4). The end of the anchor root is surrounded by a hood-like structure that is probably composed of connective tissue. The anchor point has a fine ridge or indentation running on both sides from the apex to about half way down its length towards the anchor shaft. Many, but not all, Gyrodactylus species possess a flat thin process on the ventral side of the anchor, called the anchor fold. The ventral bar processes are situated under these folds under natural conditions. On the dorsal side of the anchor, roughly opposite the anchor fold, between the anchor root and anchor shaft, is another structure where the dorsal bar attaches to the anchor (Malmberg, 1970). The relative positions of the anchor fold and the dorsal bar attachment point can be of diagnostic use for some species. Measurements are taken of the lengths of the anchor root (l.a.r.), anchor shaft (l.a.s.), and anchor point (l.a.p.) (Fig. 5a). The overall length of the anchor (l.a.) is also measured and the angle/curvature between the anchor shaft and point (Fig. 5b) is also noted. The ratio of length of anchor root to length of anchor shaft is of taxonomic significance (see Malmberg, 1970). The direction in which the anchor roots are orientated can also be useful (Fig. 5b), but variation can occur within a species. Mo (1991a) describes variation in the anchors (and marginal hooks and ventral bar) of G. salaris specimens taken from Atlantic salmon parr at different times of the year. The anchor from G. salaris, shown in Figure 5b, where its root curves slightly outwards, is typical of G. salaris taken from salmon. However, the anchor roots of G. salaris taken from rainbow trout can be straighter and rounder at the top (Mo, T.A. unpublished observations). The anchor roots of Gyrodactylus pungitii have been found to vary in orientation with respect to water temperature, being straight (and longer) during cold periods, and curved inwards (and shorter) during warm periods (Malmberg, 1970). Some anchor roots are very flexible and their shape and orientation can be altered during fixation or preservation and slide preparation. Figure 6 shows an extreme case of distortion in an anchor root from a species of the subgenus G. (Paranephrotus), possibly Gyrodactylus unicopula Gluchova, 1955 (Malmberg, 1964). G. derjavini also has very flexible anchor roots. The anchor roots of G. derjavini are usually straight, but because the roots are hollow and the root walls are quite thin, the roots can bend during preparation. If the parasite is fixed or preserved before preparation, the observation of a bent or curved


21

(usually inwards, as in Fig. 5b) anchor root is very common (T.A. Mo, unpublished observations).

Figure 6. Natural form of anchor from Gyrodactylus unicopula Gluchova 1955 is shown on the left above. The anchor root of this species is very flexible and is often found folded over on itself, as shown on the right above, following slide preparation (Malmberg, 1964)

anchor root


22

Ventral Bar The ventral bar is composed of three parts; the bar proper, two processes, which may fasten the bar to the anchors, and the ventral bar membrane (sometimes referred to as the ventral bar shield). The bar consists of three strata, dorso-ventrally positioned. On the ventral surface is a rectangular layer, usually transversely striated towards the ventral bar membrane. Its width usually corresponds to the basal width of this membrane. The middle layer extends to the ends of the bar, and is divided into two parts. The third layer, on the dorsal surface, may be totally covered by the other layers when viewed from the ventral side. However, in most species examined, a small part of it; the lateral margin, on the side nearest to the dorsal bar, is visible. It is thought that the two ventral bar processes and the ventral bar membrane emanate from this layer (Malmberg, 1970). It is often difficult to see these layers under light microscopy. Measurement points for the ventral bar are given in Figure 5d. The length of the ventral bar processes (l.p.v.b.) are measured from the posterior point where they meet the bar as this is more distinct than the anterior point. The maximum distance between the processes is measured (m.d.p.v.b.). The width of the basal part of the bar (b.w.v.b.) is measured and compared with that of the median part (m.w.v.b.). In most species these parts of the bar show distinct differences (Malmberg, 1970). The bar proper can vary greatly in size between species. The ventral bar processes can also vary in size between different species. They can be large, small or absent. The angle of orientation of the processes in relation to the median part of the bar can also be used in species identification (Fig. 5e) (Malmberg, 1970). However, within a given Gyrodactylus species, there can be large variations in size and the orientation of processes both within and between different populations of the parasite species. Variation in the size of the lateral processes has been found in Gyrodactylus specimens collected at different temperatures. G. salaris specimens collected during warm temperatures often lacked one or both of the processes, while specimens collected during cold temperatures always possessed both processes (Mo, 1991a). The ventral bar membrane can vary considerably in shape between some species, but in other cases the differences can be small and difficult to detect. The membrane is often difficult to see in preparations and is best observed under phase contrast microscopy. Most species possess a ventral bar membrane, though there are exceptions to this, eg Gyrodactylus emembranatus (Malmberg, 1970). It is also easily distorted during preparation, so determining its true shape can be difficult. It may be smooth or longitudinally ridged. The ventral bar membrane is measured as shown in Figure 5d. The ventral bar, as a whole, can display great variation in size between specimens of the same species, associated with water temperature at time of collection, but the shape remains relatively constant. Dorsal Bar Dorsal bars are normally composed of two attachment regions, for attachment to the anchors, joined by a connective structure. However, some species lack this structure (Malmberg, 1964). Studies by Malmberg (1970) have shown that the overall form of the dorsal bar is of taxonomic significance but not the detailed measurements. Therefore, only the length and width are measured. Where the bar is bent during specimen preparation, the bar is measured in two (or more) stages, to the outermost edges of the attachments. The attachments of different species can be of different size and shape, and the connective structure of different length and width. The middle of the median (connective structure) part


23

of the dorsal bar can vary greatly in shape between different species (Fig. 5f) (Malmberg, 1970). In some cases this part can also show intraspecific variation, eg presence or absence of notches. The width of the dorsal bar is measured at this point (m.w.d.b.) (Fig. 5f). The dorsal bar is seldom useful for identifying Gyrodactylus to species level, but it can give information about subgenera or species group. Marginal Hooks The marginal hook is of significant taxonomic value. Of all the opisthaptoral hardparts described here, it offers the best possibilities for distinguishing between Gyrodactylus parasites at the species level. Gyrodactylus species have 16 marginal hooks. They are divided into eight pairs and numbered 1-8 on each side of the opisthaptor (Fig. 2b). The marginal hook is composed of the sickle proper, a handle or shaft, one or two sickle membranes and a sickle-filament loop (Fig. 4). The handle articulates with the sickle. A groove is present on either side of the marginal hook sickle of at least some species of Gyrodactylus. Its presence has not been confirmed for all species (Mo, 1993; Mo and Appleby, 1990). The length of the hook proper (l.si.) and proximal (w.p.s.) and distal widths (w.d.s.) (Fig. 5a) are measured (Malmberg, 1970). In most species examined, the length (l.h.) (Fig. 5a) of the marginal hook handle of anterior marginal hooks 1, 2 or 3 (those closest to the body proper), is shorter than the posterior marginal hooks 6, 7 or 8. The only known exceptions to this are Gyrodactylus katharineri Malmberg, 1964 and G. pungitii Malmberg, 1964 (Malmberg, 1970). Therefore, it may be necessary to standardise the hook pair from which the handle is measured, for comparison between different species. Marginal hooks can vary greatly in size between different species. Details of the shape of the structures should be carefully observed. Within the marginal hooks, these details include the position of the sickle tip relative to the toe tip, the position of the heel base relative to the toe tip, size of heel, level of toe and heel in relation to handle insertion, and the sickle curve, among others (Fig. 5c). The shape of the marginal hook proper is usually species-specific in Gyrodactylus parasites. The handle shows some small differences between species. The end articulating with the sickle is usually narrower, while the other end can have a structure for attachment to the muscle. This structure is indistinct or absent in some species eg Gyrodactylus lucii. The shape of the sickle-filament loop does not vary greatly in Gyrodactylus, but its length (l.si..f., Fig. 5a) can differ. However, artificial differences in length can easily arise during specimen preparation. The number of sickle membranes (one or two) can also be used in species discrimination. The level of intraspecific variation in shape of the marginal hooks is very low. Size, as for the anchors and ventral bar, can vary with water temperature (Malmberg, 1970).


24

Cirrus The cirrus is not suitable for differentiating between Gyrodactylus species. It may not be developed in all individuals examined. Differences in size of the large spine and small spines of the cirrus seldom vary enough between species to be of taxonomic interest. However, the actual number of arched rows of small cirrus spines is sometimes useful (Malmberg, 1970). Host Gyrodactylus parasites are generally thought to be very host specific. As a result, knowing the host from which the parasite was taken can sometimes help with identification. However, it is now known that some Gyrodactylus parasites can survive and/or reproduce on more than one host species. The host range of G. salaris is large and the best studied so far (Bakke et al., 1991a; 1991b; 1991c; 1992a; 1992b; 1996; 1999). Morphological Analysis of Gyrodactylus Species Commonly Found on Salmonids in Europe Figures 5a-f and 7 show morphological characters described above for G. salaris, G. thymalli, G. teuchis, G. derjavini and G. truttae. It can be seen that G. thymalli, G. teuchis and G. salaris are very similar morphologically. The case of G. salaris/G. thymalli is particularly problematic as they cannot currently be readily distinguished using molecular techniques (as described below). In addition, morphological differences found between G. salaris infecting salmon, and G. thymalli infecting grayling, are reduced when G. salaris infecting rainbow trout is compared with G. thymalli (Malmberg, 1993; Mo, 1991b; 1994). G. salaris is a notifiable pathogen on List III of Community Fish Health Legislation 91/67/EEC, as stated in the introduction. Infection of Atlantic salmon parr with this parasite can result in severe mortalities. G. thymalli and G. teuchis have been found in countries with a G. salaris-free status. Therefore, being able to distinguish between these latter parasites and G. salaris is very important. G. teuchis can easily be distinguished using molecular techniques, described later. Gyrodactylus caledoniensis (a species described from wild and farmed Atlantic salmon in Scotland) and G. derjavini are also similar morphologically (Shinn et al., 1995). These species and G. truttae do not usually cause problems with respect to fish health, though gyrodactylosis involving G. derjavini has been described in some eastern countries (Ergens, 1983). G. derjavini and G. truttae are commonly found on salmonid fish and can be easily distinguished from G. salaris. A descriptive comparison of some of the features seen in the different Gyrodactylus species in Figure 7 is given in Table 1 and measurements in Table 2. Again, it must be emphasised that these observations come from a number of different authors and as such, caution must be exercised when using these criteria for differentiating Gyrodactylus species.

TABLE 1 A.S: Atlantic Salmon. 1 Cunningham et al. (2001); 2 Shinn et al. (1995); 3 Lautraite et al. (1999); 4 Ergens (1983) ; McHugh et al. (2000), *The diagnostic usefulness of this difference in the perceived “sharpness” of the drop (Fig. 5e) from the processes to the median part of the ventral bar, between G. derjavini and G. truttae, is debatable. In the opinion of the authors, differences in the ventral bars of G. derjavini and G. truttae are small and probably negligible.

Character G. salaris G. thymalli G. teuchis G. derjavini G. caledoniensis G. truttae Marginal hook larger than

G. derjavini/ G. truttae

very robust2 similar to G. salaris 2

heel weakly

pronounced 2 Pronounced 2 pronounced2 weakly pronounced2 pronounced2

more rounded than G. salaris 3

rounded2 less rounded2 less rounded2

base narrow Narrower than G. salaris (from A.S.)5

deep2 less deep than that of G. derjavini 2

narrow2

base is below handle insertion point

base is level with handle insertion point2

arches in centre3

arches in centre2 flat 2

toe narrow Narrower than

G. salaris (from A.S.), angular5

triangular in shape 3

very triangular very slender and angular in shape2

toe in line with heel base 2/3

25

Character G. salaris G. thymalli G. teuchis G. derjavini G. caledoniensis G. truttae toe drops

below attachment point of handle with marginal hook 3

toe base in line with attachment point of handle2

toe drops below attachment point of handle with marginal hook2

sickle point curved more slender than G. salaris (from A.S.) 5

longer than in G. salaris 1, tapering well beyond the toe 3

broad 2 slender2 slender2

point beyond the level of toe, but not as far as G. truttae2

point stops in line with or just beyond the toe2

point in line with toe 2 point tapers well beyond toe2

sickle shaft slender2 more slender

than G. salaris (from AS.)5

shorter than in G. salaris 1

broad 2 more slender than that of G. derjavini 2

slender2

curve of shaft is “broken” by a small angle in G. thymalli, whereas the sickle shaft of G. salaris and G. teuchis is constantly curved. This angle can be difficult to see in G. thymalli 1

26

Character G. salaris G. thymalli G. teuchis G. derjavini G. caledoniensis G. truttae sickle blade longer and

more curved than in G. salaris

sickle width distance

between proximal and distil widths smaller than that of G. teuchis 1

distance between proximal and distil width about 2µm (proximal part wider)1

inner aperture Oval3 rounded

compared to G. salaris/G. teuchis

rounded, more closed compared to G. salaris 3

rounded compared to G. caledoniensis 2

more square compared to G. derjavini 2

slightly larger than that of G. salaris and G. teuchis.1

Ventral bar

process rectangular1

constant and short 3/small1

similar to G. derjavini 2

more pointed and elongated than G. truttae 2

less pointed and elongated than G. caledoniensis 2

27

Character G. salaris G. thymalli G. teuchis G. derjavini G. caledoniensis G. truttae very variable2

transverse depression between processes (often seen) gives stepped appearance2 *

less stepped, but can occasionally appear stepped.

attachment opposite lower

third of dorsal bar attachment point2

opposite lower third of dorsal bar attachment point

opposite mid-point of dorsal bar attachment point3



transverse depression between processes, giving stepped appearance, seldom seen2 *

medial ridge more noticeable

than in G. truttae2

less noticeable than in G. derjavini 2

median point width

wider compared to G. derjavini 2

membrane long 3/and

tongue shaped1/3

more pointed and elongate than G. truttae2

rounder and shorter compared with G. derjavini2

Anchors similar to G.

salaris1 indistinguishable from G. truttae/G. caledoniensis 2

indistinguishable from G. truttae/G. derjavini 2

indistinguishable from G. derjavini 4/G. caledoniensis 2

larger than those of G. derjavini and G. truttae2

slightly larger than G. salaris/ G. teuchis 1

28

TABLE 2A Measurements (µm) of opisthaptoral features, used in species identification, from G. salaris, G. thymalli and G. teuchis in light microscope based studies. N: Number of parasites measured. 1(Cunningham et al., 2001 (summary of measurements taken from Mo 1991a,b,c)); 2(Shinn et al., 1995); 3(Lautraite et al., 1999); 4(Denham and Long, 1999); 5(Ergens, 1983)

G. salaris 1 G. thymalli 4, 5 G. teuchis 1, 3 Character

N Range N Range N Range

Marginal Hooks

Total length (lmh) 783 33.0- 46.5 9 37.0- 49.4 98 33.0- 39.3

Length of handle (lh) 807 26.0- 38.5 9 35.2- 39.5 99 26.7- 32.2

Length of sickle (lsi) 849 7.0- 9.5 11 7.0- 10.4 112 7.0- 9.3

Width of distal sickle (wds) 11 6.5- 7.4 21 7.0 - 7.5

Width of proximal sickle (wps) 11 5.4- 6.3 21 5.0- 5.5

Anchor

Total length (la) 872 58.0- 85.0 9 75.0- 105.0 112 60.5- 74.2

Length of anchor shaft (las) 872 43.0- 64.0 11 59.2- 77.8 114 43.6- 56.2

Length of anchor point (lap) 878 28.0- 44.0 11 33.0- 48.1 114 31.1- 40.9

Length of anchor root (lar) 875 15.0- 32.0 11 23.0- 31.6 111 17.4- 27.0

Ventral Bar

Maximal distance between processes of ventral bar (mdpvb) 708 18.5- 33.0 101 22.1- 33.0

Length of ventral bar (lvb) 753 19.5- 32.0 7 22.1-34.1 103 25.0- 34.0

Total width of ventral bar (twvb) 715 20.5- 36.5 91 26.0- 34.9

Basal width of ventral bar (bwvb) 776 7.0- 18.5 99 9.8-15.8

29

G. salaris 1 G. thymalli 4, 5 G. teuchis 1, 3 Character

N Range N Range N Range

Total median width of ventral bar (tmwvb) 719 17.0- 35.5 86 22.9- 30.3

Median width of ventral bar (mwvb) 778 5.0- 15.5 7 8.4-12.0 98 7.1-11.4

Length of ventral bar membrane (lvbm) 737 12.5- 23.0 7 18.0- 31.8 89 13.9- 21.8

Width of ventral bar membrane (attachment) (wvbm) 8 17.0- 20.5

Length of ventral bar processes (lpvb) 9 1.5- 3.0

Dorsal Bar

Length of dorsal bar (ldb) 7 20.0-30.5 7 24.0- 34.0

Median width of dorsal bar (mwdb) 7 2.0- 4.0 9 2.0- 3.5

30

TABLE 2B Measurements (µm) of opisthaptoral features, used in species identification, obtained from G. derjavini, G. caledoniensis and G. truttae, in light microscope based studies. N: Number of parasites measured. 2(Shinn et al., 1995), 6(Mo, 1993).

G. derjavini2, 6 G. truttae2 Character

N Range N Range

Marginal Hooks

Total length (lmh) 419 25.0- 38.0 117 26.3-34.4

Length of handle (lh) 421 20.0- 31.5 117 20.6-27.8

Length of sickle (lsi) 420 6.0- 7.5 117 5.6-7.5

Width of distal sickle (wds) 168 3.8-5.6 117 4.1-5.6

Width of proximal sickle (wps) 168 4.1-5.6 117 4.4-5.6

Hamulus

Total length (la) 421 46.9-65.6 117 50.0- 66.3

Length of anchor shaft (las) 412 32.5-48.1 117 35.9- 48.8

Length of anchor point (lap) 420 23.8- 35.0 117 26.3- 38.1

Length of anchor root (lar) 412 12.5-23.1 117 8.8- 21.3

Ventral Bar

Maximal distance between processes of ventral bar (mdpvb) 244 23.5- 31.5

Length of ventral bar (lvb)

Total width of ventral bar (twvb)

Basal width of ventral bar (bwvb)

Total median width of ventral bar (tmwvb) 244 17.0- 25.0

31

G. derjavini2, 6 G. truttae2 Character

N Range N Range

Median width of ventral bar (mwvb) 413 4.4 –10.0 117 5.3- 9.4

Length of ventral bar membrane (lvbm) 392 9.4-18.2 117 9.4- 18.8

Width of ventral bar membrane (attachment) (wvbm)

Length of ventral bar processes (lpvb)

Dorsal Bar

Length of dorsal bar (ldb) 117 21.3- 30.6

Median width of dorsal bar (mwdb) 117 1.3-3.1

32


33

Anchor

G.s G.teu G. d G. tru.G.thy

Marginal Hook

G. s G. tru.G. thy G. teu G. d

Ventral Bar

Figure 7. Opisthaptoral hardparts used in identification of Gyrodactylus species. Figures reproduced from 1Ergens (1983) ; 2Cunningham et al, (2001); 3Mo (1993). * Modified figure of opisthaptor taken from Lautraite et al., (1999). One division = 10µm.

Opisthaptor

G.sG. thy G. d G. tru.G. teu

1 1 1* 1

2 22 1 1

2 1111

G.s G.teuG.thy G.d G. tru.

2 2 3 12

G. thy G. s G. tru.G. teu G. d

1 1 2 1 1


34

Gyrodactylus 'Quick' Identification Checklist The principal point of interest for the majority of fish health laboratories is whether or not G. salaris is present. Therefore, where expertise in Gyrodactylus parasite identification is not available, or where time is limited, being able to classify a specimen as not being G. salaris, without actually identifying the parasite to species level, can be sufficient. For this purpose, the following checklist has been assembled. This checklist concentrates on easily recognisable morphological features seen under the light microscope, with a minimum number of measurements required. Where a specimen cannot be classified as “not G. salaris”, then it must be examined in more detail. If the specimen cannot be identified, it may be classified as a “no identification” with respect to morphological identification and may be processed using molecular techniques (as are those specimens determined as “not G. salaris” for the purpose of further confirmation). Division of the specimen into the opisthaptor, which can be retained for further morphological examination, and the body, which can be used in molecular diagnostic techniques, is preferable in the case of specimens that cannot be identified from morphology alone. The opisthaptor may be prepared as described for whole mounts. This checklist is based mainly on distinguishing G. salaris from G. derjavini and G. truttae, while attempting to distinguish G. derjavini from G. truttae. G. derjavini and G. truttae are found commonly on salmonids in Europe. However, it is important to remember that there are other species which might be present, eg G. thymalli and G. teuchis (which are similar in appearance to G. salaris), G. caledoniensis, and potentially new and as yet uncharacterised species.


35

Gyrodactylus “Quick” Identification Checklist

1. Anchors Comments

a) Is the anchor fold on the ventral bar opposite the midpoint of the dorsal bar attachment?

If yes, then it is not G. salaris.

b) Are the roots obviously curved inwards? If yes, then it is probably not G. salaris

c) Are the roots very straight with little curvature?

G. derjavini, G. truttae and G. salaris can all display straight anchor roots, but in our experience, the anchor roots of preserved G. derjavini specimens are often pointing inwards in slide preparations.

d) Do the roots angle outwards? This is often seen in G. salaris.

e) Is the total length of the anchor greater than 70 µm?

This indicates it is not G. derjavini/G. truttae The length of the anchors of G. derjavini and G. truttae is usually less than 70µm. However, G. salaris also can have anchors less than 70µm in length.

2. Ventral Bar Comments

a) Are the ventral bar processes large (wing shaped)?

If yes, then it is not G. salaris.

b) Are the processes small, and does the overall structure of the processes and ventral bar together give an “even” appearance anterio-posteriorly (rectangular)?

This is a G. salaris trait.

3. Marginal Hooks (of adult &/or embryo) Comments

a) Is the hook quite thick and robust ? If yes, possibly G. derjavini

b) Is the base deep and does it have a pronounced heel?

If yes, possibly G. derjavini.

c) Does the tip of the base (toe) obviously point downwards below the bottom of the base/heel?

If yes, possibly G. truttae.

d) Is the sickle point in line with, or just beyond the front (toe) of the base?


e) Is the tip of the sickle obviously extending beyond the front (toe) of the base?

If yes, possibly G. truttae/G. salaris.

f) Is the sickle point quite thick and curved downwards?


g) Is the sickle point quite straight and slender and only slightly curved downwards?

If yes, possibly G. truttae.

h) Is the curve of the sickle slender and curving downwards?

This is a G. salaris trait.


36

Diagnosis Using Molecular Means (see Cunningham (1997); Cunningham et al., (1995a and b)). Molecular tests developed for the diagnosis of Gyrodactylus parasites have targeted regions of the ribosomal RNA (rRNA) gene array, also known as ribosomal DNA, or rDNA. This array consists of three genes for ribosomal RNA; the 28S large subunit, the 18S small subunit, and the 5.8S, interspersed with spacers. The internal transcribed spacer regions (ITS1 and ITS2) and the external transcribed spacer (ETS) at the 5' end of the 18S gene are transcribed, but the intergenic spacer (IGS) is not. These arrays occur in tandem repeats throughout the genome and as such provide a high number of targets for amplification by polymerase chain reaction (PCR). Figure 8 shows a diagrammatic representation of a ribosomal gene array. PCR is employed to amplify certain parts of the ribosomal RNA genes or spacers for further examination. The power of PCR to amplify DNA from very small amounts of starting material enables several different amplification reactions to be carried out from an individual parasite, and thus several different regions of the genome can potentially be examined.

Figure 8. Ribosomal gene array, showing positioning of genes and intervening spacers.

Internal transcribed spacer(ITS)

118S 5.8S 28S 18S28S

Intergenic spacer(IGS)

2

External transcribed spacer (ETS)

Genes


37

Preparation of Samples

• Remove individual specimens from live or preserved fish. • Wash or blot preserved parasites to remove excess liquid. • Place a single parasite in a 0.5 ml microfuge tube, containing 7.5 µl lysis buffer

(0.45% IGEPAL, 0.45% Tween 20, 60 µg/ml Proteinase K). • Incubate the tube for a minimum of 20 minutes at 65ºC to allow digestion of parasite

tissue and release of DNA. • Incubate the tubes at 95ºC for 10 minutes to inactivate the Proteinase K. • Check lysate in tube, under a dissecting microscope, to see if parasite has been

digested. If not, a second aliquot of Proteinase K can be added and the tube left to digest at 65ºC for longer. Check intermittently to see if parasite has been lysed.

• The tubes must again be incubated at 95ºC for 10 minutes to inactivate the Proteinase K.

This lysate is used as DNA template in PCR reactions without further purification. The V4 Region of the Small Subunit Ribosomal RNA Gene (see Cunningham et al., 1995a; 1995b) The V4 region is one of a number of variable regions found within the 18S small subunit ribosomal RNA gene. It is often used in the identification of species by molecular means. PCR amplification of the V4 region • Make up a PCR mix to contain the following reagents in each reaction tube; 1x PCR

buffer, 1.75 mM MgCl2, 200 µM dNTPs, 1 µM of each primer (V4F: 5'-CTA-TTG-GAG-GGC-AGT-CT-3' and V4R: 5'-CTT-TTC-AGG-CTT-CAA-GG-3'), dH2O to a final volume of 16.5 µl. Prepare sufficient mix for each specimen, one negative control, and one extra tube to compensate for inaccuracies in pipetting.

• Aliquot 16.5 µl of the mix into individual, labelled microfuge tubes and overlay with mineral oil.

• Add 2.5 µl of parasite lysate as prepared above to individual tubes. • Add 2.5 µl of dH2O to the negative control. • Incubate the tubes at 95ºC for five minutes. • Add one unit Taq polymerase (in a 1 µl volume) to each tube, while still at 95ºC. • Subject the PCR tubes to 30 cycles of 92ºC for one minute, 50ºC for 30 seconds and

72ºC for 30 seconds, followed by one cycle of 72ºC for five minutes. Visualisation of PCR products on gel

• Run 4 µl of the above PCR product on a 2% (w/v) agarose gel containing ethidium bromide. Warning: Ethidium Bromide is a potential mutagen and suitable safety measures should be observed when handling it, or gels and solutions in which it is present.

• Run a suitable DNA size marker (eg 100 bp ladder) and a mass marker alongside the V4 PCR product.

• Determine the size and concentration of the V4 PCR product from the gel; a 358bp product should be present.

• Store the remaining PCR products at 4ºC. If this PCR product is to be analysed using hybridisation with DNA probes, it is best applied to membranes within two days of PCR amplification.


38

Restriction digest of the V4 region Restriction enzyme recognition sites that allow discrimination of G. salaris, G. derjavini and G. truttae have been identified in the V4 sequence (Cunningham et al., 1995a). If the V4 region is amplified at high concentration, no non-specific amplification products are formed and PCR primers and excess dNTPs do not appear at high concentrations in the agarose gel, it is possible to carry out diagnosis following digestion of PCR product with combinations of two enzymes, such as DdeI and HaeIII. However, in practice this is not always straightforward; PCR products may be at low concentration and thus restriction fragments difficult to visualise and the restriction fragments are relatively small and require gels with high concentrations of agarose for clear separation. Therefore, this method is not described in detail here, but can be found in Cunningham et al. (1995a). Generation of probes Probe hybridisation as a diagnostic method is based on the principle that short species-specific sequences will only hybridise to complementary DNA, when conditions for the hybridisation reaction are sufficiently stringent. Therefore, they should only bind to the species from whose DNA the probe was designed. Figure 9 shows a diagrammatic overview of the technique.


39

Figure 9. Amplification of the V4 region of ribosomal DNA followed by probe hybridisation, as a diagnostic method for Gyrodactylus. Oligonucleotides corresponding to the V4 variable regions of G. salaris (GsV4B), G. derjavini (GdV4) and G. truttae (GtV4) have been designed (Fig. 10) (Cunningham et al., 1995b). The probes have been used to successfully separate G. salaris from G. truttae and G. derjavini. However, the V4 region of G. salaris and G. thymalli is identical and, as a result, cannot be used to differentiate between these two species. The probes described by Cunningham et al. (1995a and b) also cannot differentiate G. salaris and G. teuchis as the G. salaris probe was designed within a region of V4 sequence common to both species (see

Confirm presence of V4 PCR product on gel

Hybridise labelled probe generated from different control species, to separate membranes

V4 labelled probe hybridisation to specimen DNA

Detect bound labelled probes

Wash unbound labelled probes

Bind specimenV4 PCR product

to eachmembrane

Bind control V4 DNA

to each membrane

Lysis buffer

PCR

V4

PCR mix

Prehybridise prepared membrane


40

Cunningham et al., 2001). Other parts of the V4 region of G. teuchis differ significantly from that of G. salaris, G. truttae, and G. derjavini and therefore new probes to differentiate all four species could easily be designed. This has not as yet been done, as other molecular methods (see ITS-RFLP below) are currently being used for successful differentiation of these parasites. An important consideration to be taken into account when using the hybridisation method below with new probes, is the dissociation temperature of the probes. This may change the hybridisation temperature and the stringency of the washing steps. These values must be optimised experimentally. Sequences of the V4 region of the small subunit (18S) ribosomal RNA gene, and location of primers and probes are given in Figure 10. G.salaris CTATTGGAGGGCAGTCTGGTGCCAGCAGCCGCGGTAACTCCAGCTCCAATAGCATATAT 59 G.teuchis ........................................................... G.derjavini ........................................................... G.truttae ........................................................... G.salaris TAAAATTGCTGCAGTTAAAAAGCTNCGTAGTTGGATCTGGGTTCTGGTTTGGAGACTGCT 119 G.teuchis ............................................................ G.derjavini ............................................................ G.truttae ............................................................ G.salaris TGCTCTTAGTGAATTGATTTCATGAAGCTTTGGGCAGCGGTACTTCTAGGCCGAATCTTC 179 G.teuchis ............................................................ G.derjavini ...........G....T........................................... G.truttae ..T..A.....G....T..................…........................ G.salaris CAGCTGTGTCTGCATAAGGCTTCGGCTTTGTGTAGATAGATTCGTTGTATGTTAGTTCCC 239 G.teuchis ................................................G...A....... G.derjavini ................G..T......C................................. G.truttae ................................................G.TGA....... G.salaris TCACGGGTCTACTTCTTCGTTTCTATACGCTGTAATGCCTTTAATCGGGTGTTCAGTGTG 299 G.teuchis ..........T.AC.............................................. G.derjavini ............................................................ G.truttae ..........T.ACT....GA....................................... G.salaris GACAGCACGTTTACTTTGAACAAATTTGAGTGCTCAAAGCAGGCCTTGAAGCCTGAAAAG 359 G.teuchis ............................................................ G.derjavini ............................................................ G.truttae ............................................................ Figure 10. Sequence alignment of the V4 region of the 18S ribosomal RNA gene from G. salaris, G. teuchis, G. derjavini and G. truttae. The PCR product generated by the V4 primers extends beyond the V4 region at both 5' and 3' ends. The V4 region is highlighted in yellow. V4 PCR primer positions are highlighted in grey. Boxed sequences indicate oligonucleotide probes GsV4B(679-694), GdV4 (747-761) and GtV4 (797-811). (•) base identical to G. salaris; (N) base unique to one species.


41

The method described in the following section uses the probes GsV4B, GdV4 and GtV4. The V4 oligonucleotide probes were synthesised commercially. Probe labelling was carried out according to the protocols in the DIG System User’s Guide using DIG labelling kits (Boehringer Mannheim). These kits use chemiluminescent detection of bound probes and therefore are safer than radioactively labelled probes. • Label probes GsV4B (5'-GTG-AAT-TGA-TTT-CATG-3'), GdV4 (5'-GGG-TTT-CGG-

CCT-TGT-3') and GtV4 (5'-GTC-TTC-ACT-TTC-GGA-3') with dioxigenin-11-ddUTP using the reagents and protocol supplied with DIG Oligonucleotide 3'-end Labelling Kit.

• Estimate the yield of labelled oligonucleotide, as described in the manufacturer’s protocol, by comparing the signal intensities with those produced by labelled control DNA (provided with Kit).

Preparation of membranes for hybridisation Control V4 PCR products amplified from known samples of, or cloned V4 DNA from, G. salaris, G. derjavini and G. truttae should be included on all membranes.

• Mark positively charged nylon membranes with a pencil to indicate position of

samples to be tested, and controls (Figure 11).

Figure 11. Example of a marked membrane, for 20 samples (1-20) and G. salaris (G.s.), G. derjavini (G.d.) and G. truttae (G. t.) controls.

• Denature the sample and control DNA by heating 10 µl aliquots of each PCR product

in 0.5 ml microfuge tubes in a boiling water bath or a dry block heater at 99ºC for 10 minutes.

• Place tubes directly on ice. • Immediately spot 1 µl of each solution onto each of three positively charged nylon

membranes (one for each probe being used) at the appropriate mark for that sample. • Bake membranes at 120ºC for 30 minutes.

G.s

G.d

G.t

1 8

10 16

17 20


42

• Store membranes at room temperature until hybridisation is carried out. • Store the remainder of the PCR products from the diagnostic samples at 4ºC until

diagnosis has been made. Hybridisation of labelled probes to membrane-bound samples • Place replicate membranes in separate hybridisation bottles or trays and incubate in

20 ml prehybridisation buffer (as per DIG kit) in a rotating chamber, or on an orbital shaker, for one hour at 30ºC.

• Dilute labelled oligonucleotides separately in hybridisation buffer (as per DIG kit) to a concentration of 10 pmol/ml.

• Discard prehybridisation buffer and replace with 6ml of the hybridisation buffer containing the probe (10 pmol/ml), one probe type per membrane.

• Incubate membranes in rotating chamber, or on an orbital shaker, at 30˚C for 3.5 hours.

• Following hybridisation, decant probe solutions into separate tubes and store at –20˚C for re-use.

• Wash membranes twice at room temperature for 5 minutes each wash in 2 x SSC (standard saline citrate; 0.3M sodium chloride (NaCl), 0.03M sodium citrate (C6H5Na3O7), pH7.0; 0.1% (w/v) SDS (sodium dodecyl sulfate)). SSC can be purchased as a 20x ready made stock solution.

• Wash membranes twice at 30˚C for 15 minutes each wash in 0.1 x SSC, 0.1% (w/v) SDS.

Detection of labelled probes bound to membrane Chemiluminescent detection of bound probe is carried out using reagents and the protocol as given in DIG Chemiluminescent Detection Kit and Wash and Block Buffer Set. Analysis Species are identified by comparing the signals from diagnostic samples with signals from control samples of G. salaris, G. derjavini, and G. truttae. Figure 12 shows a developed film following exposure to membranes containing bound Gyrodactylus V4 PCR products hybridised with DIG-labelled V4 probes.


43

Film

Imprint of membranes with: • marks indicating position of • Gyrodactylus DNA samples and • control V4 PCR product • membrane hybridised with G. salaris • V4 probe (G. salaris control DNA • detected) • membrane hybridised with G. derjavini • V4 probe (G. derjavini control DNA and • sample DNA detected) • membrane hybridised with G. truttae • V4 probe (G. truttae control DNA • detected)

V4 PCR samples bind G. derjavini V4 probe, but not G. salaris/G. truttae V4 probes.

nonspecific background staining

Figure 12. Diagnosis of Gyrodactylus species using hybridisation of DIG-labelled V4 probes from known Gyrodactylus species, to V4 PCR products from unidentified Gyrodactylus parasites. s; Gyrodactylus salaris, d; G. derjavini, t; G. truttae.


44

Ribosomal RNA internal transcribed spacer (ITS) (see Cunningham (1997)) The internal transcribed spacer (ITS) region of ribosomal DNA lies between the 18S small subunit ribosomal RNA gene and the 28S large subunit ribosomal RNA gene. It is composed of two regions, the ITS1 and ITS2, which are separated by the 5.8S ribosomal RNA gene. The spacers are transcribed but do not form part of the mature ribosome and as such are thought to be under less conservation pressure than coding regions and so acquire a greater number of mutations between species. The ITS has been widely used in the development of molecular diagnostic techniques for species discrimination, particularly in nematode and trematode parasites. Restriction fragment length polymorphism (RFLP) uses the properties of restriction enzymes that recognise specific DNA sequences to which they bind and then digest the DNA. A large number of restriction enzymes exist, recognising different DNA sequences. RFLP is a way of differentiating between different species based on the gain or loss of restriction enzyme sites, which in turn reflects differences in the DNA sequence. A diagrammatic overview of the procedure of PCR amplification of the ITS region, followed by restriction enzyme analysis, is given in Figure 13. Sequences of the ITS region of a number of Gyrodactylus parasites, showing PCR primer sites and restriction sites, are given in Figure 14.


45

Figure 13. Amplification of the ITS region of ribosomal DNA followed by restriction digest and analysis of restriction fragment sizes as a diagnostic method for Gyrodactylus.

Restriction Fragment Length Polymorphism (RFLP) analysis of specimen DNA

Confirm presence of ITS PCR product on gel

Same RFLP pattern

Specimen Controls

Restriction digest mix

ITS

PCR

PCR Mix

Lysis buffer

G.salaris TTTCCGTAG-TGAACCTGCGGAAGGATCATTAAACATCGTTTCCTTACTTTGTGGTGACT 59 G.teuchis .........G...................................-.A.........T.. 59 G.derjavini .........G.................................................. 60 G.truttae .........G...................................-.A......A..... 59 G.salaris CAGTGGTGTCATTGCCTAAAATCAAAGGATT------TTAAA------GAATTARAAGCA 107 G.teuchis ...........A.........AA.CC.A...AACT.............T..A..A...T. 111 G.derjavini ...A.T.................TGC.....TAAAAGA....TATTCAT..A..A...T. 120 G.truttae ...........A...G.......TCG...............-........GA..A...T. 106 G.salaris GTTAATGGTGATTCGTTTGTTATTGCATGGTTACGGTATAATGATAT-ACCTTGAAGAAT 166 G.teuchis .CA.T..T.C.GA....CTG..........C..A......GA..GGGT....A..G...G 171 G.derjavini .CA.T..CGT.......CAG..A-......C.TA.C..---------.------------ 157 G.truttae .CA.T....T.......CAG..A.......C..-....ACT..T..-.-....C.....G 162 G.salaris AAAGAAT-----------------AAGGGTGGTGGCGCACCTATTCTACAA--GCAGA-- 205 G.teuchis TCG..G.CA..............G..A...A...........G....GTT.GT.AG..CT 217 G.derjavini ---.---...................A...A..T............AG.T.G.T.TT... 191 G.truttae ...T.G.GATTATGAGATAGTTAG..A...A..A........G......T.TA.AC.... 220 G.salaris -------A--CTGGTTAATAA----------------GATC------------------- 221 G.teuchis CCTAGGA.GT.A.AG...G..AGGTAGTGGCGCACCT.T..TGTTAGTGAGGACTTCAAG 277 G.derjavini ........TA.A.ACA.GAG.TGA..............T..................... 212 G.truttae ........GT....GGC.C..CGAAGGAGAAAC........................... 250 G.salaris -----GATTCCGA-GTGA---CGATCGTGGGGCAAAAT--AAATCCAGCTTGGGGAACTG 270 G.teuchis ATAAA....GTT.A....GAA.A....A..A...C..AGT...........A........ 337 G.derjavini .....---.GA.CA.A......C...A.......CC.A..--.................. 257 G.truttae .......G....TC.A......C...........C..CG..................... 301 G.salaris GTTAACCATGGCATTATAC-GAGCAAGATGATTCCGAACGAGATTCTTTTAACATAGCAA 329 G.teuchis ....GT..AA......C.....A..---................A..............T 393 G.derjavini ..-.GT......G...C..C..A....-A...A.................T...A..... 315 G.truttae ..-.GT...C....A.C..-..A.....GT..A..........................T 359 G.salaris TGAACACACGCTGTTTCATGCGCAACCAATCTGCCCTAT-AAAATTGGAGAGTGATTAGA 388 G.teuchis .......T................T..............A....C............... 453 G.derjavini ...-..T................................A...C.........A...... 374 G.truttae ...T....................TA............-..................... 417 G.salaris TTGCTCACCCACCGTCGTTTAGATGGTTGACATTAAAACG-CTCATTGGAGTGAACTGGT 447 G.teuchis ...............................T..G...TTA................... 513 G.derjavini ...............Y................A......TA................... 434 G.truttae ......................................ATA................... 477 G.salaris AGTCTTCCGAGCTAAAATGGTAATGGCTAGTCTCGGTAAGGTCTGACTATCGGTTCGGCT 507 G.teuchis ..................T......................................... 573 G.derjavini ........................A................................... 494 G.truttae ....................................A....................... 537 G.salaris ACGGCCAGCTCAATGTAGTATCCGCTATTACCGAAACA--TACACTACAGTGGTTCGATA 565 G.teuchis ...............................TA...GT...TAT.............T.. 631 G.derjavini ................................A...AG.T.T.T................ 553 G.truttae ....................A...............A.CT.T.T................ 597 G.salaris GAGTTCCACACTCACTGCCTCTGCACCTTCGGGTGAACAGTCCGTAGTGCTTAGCGCCCC 625 G.teuchis .............G......................T...AAA................. 691 G.derjavini ....................................C...AA.................. 613 G.truttae .............G......................C...AA.................. 657

G.salaris GTCAAAAGGGAAGAAGCTTTGGTTTATTACAACTCCATGTGGTGGATCACTCGGCTCACG 685 G.teuchis ..T......................................................... 751 G.derjavini T.A...C............A.C...................................... 673 G.truttae ............................................................ 717 G.salaris TGACGATGAAGAGTGCAGCAAACTGTGTTAACCAATGTGAAACGCAAACTGCTTCGATCA 745 G.teuchis ............................................................ 811 G.derjavini ............................................................ 733 G.truttae ............................................................ 777 G.salaris TCGGTCTCTCGAACGCAAATGGCGGCTAAGGGCTTGCTCTTAGCCACGTTCGATCGAGTG 805 G.teuchis ............................................................ 871 G.derjavini ............................................................ 793 G.truttae ............................................................ 837 G.salaris TCGGCTTTTACCTATCGTAACGCTTAATTAGTTACGGATTGGGAAGTATACCATGGCTAT 865 G.teuchis ............................................................ 931 G.derjavini ............................................................ 853 G.truttae ............................................................ 897 G.salaris GCGATTAACTTGTTGTTGAAAGTTGAAACACGGGGTATTACACGGCCTTTACGGTTTGCC 925 G.teuchis .....................A........T..............T.............. 991 G.derjavini ...............C.T.............C............................ 913 G.truttae .....................A.....R...AC............T.............. 957 G.salaris CTGTGGTGTTCTGATTCTGGTATTACACGGACTTTACGGTTTGCTAGATGAAGTTCACAT 985 G.teuchis ..............................T....................T........ 1051 G.derjavini ..................A...........C............................C 973 G.truttae ..................A...C.......T...........A......A........GC 1017 G.salaris TCGATGAGTATGCGGCTTCTGAGTATTACACGGACTTTACGGTTTGCTCGGAAGTTAAAG 1045 G.teuchis ..........GA................................................ 1111 G.derjavini ..A....T..G......................C.......................... 1033 G.truttae ..A....T..GA.................................A.............. 1077 G.salaris ACCATTCTTTCATACACGGCCTTTACGGTTTGATAGAATGAGAAATAGCTCTAGTGGTTC 1105 G.teuchis ........G................................................... 1171 G.derjavini ..................................T.....C...T............... 1093 G.truttae ..................A.T...................C...G............... 1137 G.salaris TTCCTTAATTGCTTGGGTAGTATTGTTGTGTACTTTATGGTCTGCTCTGCACAGGGTGCG 1165 G.teuchis ..................................C......................... 1231 G.derjavini ..........A................................................. 1153 G.truttae ..........A....................G............................ 1197 G.salaris TGGCTTAGTTCGCTTTGTAACGCTGTACTGAAGTAGAGATAGATTTGTGCATGATATACC 1225 G.teuchis ..............................T...G...T..................... 1291 G.derjavini ......................................T..................... 1213 G.truttae ..............................TT..T...T......G.............. 1257 G.salaris CAGTGAAAATAAGTCCTGACCTCGATTCGAGCGTGAATACCCGCTGAACTTAAGCATA-T 1284 G.teuchis ........TA.................................................. 1350 G.derjavini ............................................................ 1272 G.truttae ...............................................R..........A. 1317 G.salaris CACTAAGCGGAGGA 1298 G.teuchis .............. 1364 G.derjavini .............. 1286 G.truttae .............. 1331

Figure 14. Sequence alignment of internal transcribed spacer (ITS) regions of the ribosomal DNA, from G. salaris, G. teuchis, G. derjavini and G. truttae. 5.8S ribosomal gene is highlighted in yellow. ITS primer positions are underlined. The 3’ end of the 18S ribosomal gene and 5’ end of the 28S ribosomal gene, which are incorporated in the ITS PCR product, are highlighted at the 5’ and 3’ ends. Hae III restriction sites are highlighted in grey throughout the sequences. (.) base identical to that of G. salaris. (-) gap in sequence

46

46


47

PCR amplification of ITS • Make up a PCR mix containing reagents in the following concentrations for each

reaction; 1x PCR buffer, 1.5 mM MgCl2, 200 µM dNTPs, 1 µM of each primer (ITS1: 5'-TTT-CCG-TAG-GTG-AAC-CT-3' and ITS2: 5'-TCC-TCC-GCT-TAG-TGA-TA -3'), dH2O to a final volume of 16.5µl for each reaction.

• Prepare sufficient mix for each specimen, one negative control, and extra mix to compensate for pipetting inaccuracies.

• Aliquot 16.5 µl of the mix into individual, labelled microfuge tubes and overlay with mineral oil.

• Add 2.5 µl of parasite lysate, prepared as above, to the appropriate tubes. • Add 2.5 µl of dH2O to the negative control. • Incubate the tubes at 95ºC for 5 minutes. • Add one unit Taq polymerase (in a 1 µl volume) to each tube, while still at 95ºC.

While Taq polymerase added to the PCR mix at this stage, in theory increases specificity, we have added Taq with the other reagents in the bulk mix and have only rarely obtained non-specific PCR products. PCR conditions may however need to be optimised, as differences in thermocyclers, DNA polymerases and other reagents, can influence specificity.

• Subject the PCR tubes to 28 cycles of 94ºC for one minute, 50ºC for one minute and 72ºC for two minutes, followed by one cycle of 72ºC for five minutes.

Visualisation of ITS PCR products on gel • Run 4 µl of the PCR product on a 1.5% (w/v) agarose gel containing ethidium

bromide Warning: Ethidium Bromide is a potential mutagen and suitable safety measures should be observed when handling it, or gels and solutions in which it is present.

• Run a suitable size marker (eg 100 bp DNA ladder) alongside the ITS PCR product. • Determine the size of the ITS PCR product from the gel; a PCR product of

approximately 1300 bp should be present. Note: A positive control should be included. Suitable positive controls include diluted ITS PCR products, generated using the primers ITS1 and ITS2, from any of the Gyrodactylus species (or cloned ITS) for which the ITS has been characterised. Absence of PCR products from sample specimens alone (when a positive control is not present), does not necessarily indicate that the PCR reaction has not worked. It may be due to sample storage or preparation prior to PCR. Restriction enzyme digest of the ITS

• Label the appropriate number of 0.5 ml microfuge tubes (eg label with date, sample

number and “digest”). Label four additional 0.5 ml microfuge tubes for the controls G.s (G. salaris), G.d (G. derjavini), G.t (G. truttae) and G.te (G. teuchis).

Note: prepare enough controls, so that controls from all four species accompany each row of diagnostic samples on the electrophoresis gel. Controls can be obtained by amplification of ITS products from species which have been reliably identified using morphological characteristics, or for which the ITS products have been confirmed by DNA sequencing. To maintain a continuous stock of controls, the ITS products can be cloned and stocks of recombinant plasmids containing ITS isolated. ITS products can then be obtained by PCR amplification, using these plasmids as template.


48

• Add 8.8 µl of ITS PCR product (approx 200 ng) from diagnostic samples, into the appropriately labelled tubes.

• Add 5 µl of control ITS products (approx 200 ng) into appropriately labelled tubes. • Add 3.8 µl of sterile distilled water to each control tube. • Make up a restriction mix allowing 1 µl of the appropriate 10x restriction buffer

(supplied with restriction enzyme), and 0.2 µl (2U) of the restriction enzyme HaeIII, per reaction, plus additional buffer and enzyme to compensate for pipetting inaccuracies.

• Mix well and briefly centrifuge to collect contents at bottom of tube. • Add 1.2 µl of this mix to each reaction tube and mix carefully by pipetting up and

down. • Centrifuge briefly to collect contents at bottom of tubes. • Incubate at 37ºC for a minimum of 1.5 hours. Reactions can be left overnight at

37ºC. Visualisation of restriction digest products on gel • Run the entire contents of the above digests on a 2% (w/v) agarose gel containing

ethidium bromide Warning: Ethidium Bromide is a potential mutagen and suitable safety measures should be observed when handling it, or gels and solutions in which it is present.

• Run a suitable size marker (eg 100 bp DNA ladder) alongside the digest products, and ensure that each row on the gel contains a full set of controls with which the fragment sizes of diagnostic samples can be compared.

• Determine the size of the restriction fragments by comparing with controls and markers.

Note: the expected sizes of DNA fragments obtained from HaeIII digestion of the ITS of G.salaris, G.derjavini, G.truttae and G.teuchis is given in Table 3. DNA fragments smaller than 50 bp will be difficult to see on an agarose gel. However, the larger fragment sizes can be used to clearly differentiate between the four species used as controls. A diagrammatic overview of the analysis of RFLP patterns obtained on restriction digest of Gyrodactylus ITS PCR product is given in Figure 15. Figure 16 shows an agarose gel, following visualisation under UV light, with undigested and digested ITS PCR fragments from Gyrodactylus parasites.


49

TABLE 3 Expected restriction fragment sizes obtained on digestion of ITS PCR product from Gyrodactylus species using the restriction enzyme Hae III.

Fragment sizes (bp) Restriction Enzyme G. salaris G. teuchis G. derjavini G. truttae

790

577

Hae III 511 553 541

498

399 400

234 234 234

154

63

46

45 Hae III recognises the sequence -GGCC- Restriction site GG|CC CC|GG


50

Figure 15. Restriction sites, fragment sizes, and expected gel pattern following digestion of the named Gyrodactylus parasites with the restriction enzyme HaeIII.

Enzyme restriction sites and fragment sizes

G.salaris

511 399 154 234

G. teuchis577 553 234

G. truttae541 790

G. derjavini498 400 23445 63 46

HaeIII restriction digest of Gyrodactylus ITS PCR products

Restriction fragment patterns on electrophoresis gel

G.d: G. derjavini

G.s G.te G.d G.tMarker G.s: G. salaris

G.te: G. teuchis

G.t: G. truttae

500bp


51

Figure 16. Agarose gel following exposure to UV light, showing undigested ITS PCR products from Gyrodactylus salaris (s), G. derjavini (d) and G. truttae (t) on the right of the gel, and digested ITS PCR products, using the restriction enzyme Sau 3A from the same parasites on the left of gel. The first lane contains a 100bp molecular mass ladder. The three species can be clearly differentiated from each other based on their respective RFLP patterns.


52

The volume and/or concentration of DNA used in the control digests can be reduced if fragments are too bright to allow clear definition of the different fragments. If the concentration of ITS PCR products in the diagnostic samples is too low to allow visualisation of the restriction fragments on the gel, then the amount of PCR product, restriction enzyme and buffer used per reaction can be doubled to give a final volume of 20 µl. The need for doubling the concentration of PCR product in the restriction digest reaction can be judged from the concentration of ITS product on the initial electrophoresis gel after amplification. 20-200 ng/µl PCR product is normally suitable. The method of ITS amplification, followed by restriction digest, is quicker to perform than V4 probe hybridisation. V4 hybridisation is useful when large numbers of samples have to be processed at the same time. Both V4 and ITS-RFLP analysis have been found to consistently agree with morphological analysis, once a Gyrodactylus species has been clearly characterised by both methods. Amplification of the ITS1 and ITS2 regions of Gyrodactylus ribosomal DNA We have found that when PCR amplification of the full ITS region, as described above, fails due to the quality of sample DNA, the amplification of the smaller ITS1 or ITS2 regions is sometimes successful. Amplification of ITS1 PCR amplification of the ITS1 region was carried out using primers ITS1A (5'-GTAACAAGGTTTCCGTAGGTG-3') which lies at the conserved 3' end of the Gyrodactylus 18S ribosomal RNA gene and upstream from the ITS1 primer site, and ITSR3A (5'-GAGCCGAGTGATCCACC-3') which binds to the conserved 5' end of the 5.8S ribosomal RNA gene. All other amplification reagents and conditions were as for the full ITS region above. Amplification of ITS2 PCR amplification of the ITS2 region was carried out using primers ITS4.5 (5'-CATCGGTCTCTCGAACG-3') which lies at the conserved 3' end of the 5.8S gene, and ITS2 (5'-TCCTCCGCTTAGTGATA-3') as used for the full ITS. All other amplification reagents and conditions were as for the full ITS region above. Restriction enzyme digest of ITS1 Analysis of the DNA sequence of the ITS1 region predicts that digestion with restriction enzymes HinfI and NIaIII will produce fragment patterns that will differentiate G. salaris, G. teuchis, G. derjavini and G. truttae. Some of the predicted fragment sizes are less than 100 bp. Therefore, agarose gels used in visualisation of the digest fragments should be of at least 2% (w/v) concentration, and the gels should be run at relatively low voltage, eg 75V, to obtain better resolution. The restriction sites for the restriction enzymes HinfI and NIaIII are given in Table 4, and predicted fragment sizes for the different Gyrodactylus species are given in Table 5.


53

TABLE 4 Restriction sites for enzymes HinfI and NIa III.

Restriction Enzyme Hinf I NIa III

Restriction Site G|ANTC CATG|

CTNA|G |GTAC

TABLE 5 Predicted Gyrodactylus ITS1 ribosomal DNA HinfI and NIa III restriction fragment sizes.

Fragment sizes (bp) Enzyme

G. salaris G. teuchis G. derjavini G. truttae

Hinf I 371 385 373 373

212 240 195

105 149 94 93

93 39 90

77

61

12

NIa III 315 317 318 317

173 223 185 244

144 177 82 172

69 50 68

41

36

18 18 18 18 Restriction enzyme digest of ITS2 Of the restriction enzymes already mentioned, only HaeIII will give restriction patterns that distinguish G. salaris, G. teuchis, G. derjavini and G. truttae. As for ITS1, some of the predicted restriction fragments are less than 100 bp and agarose gels used in visualisation of the digest fragments should be of at least 2% (w/v) concentration, and the gels should be run at low voltage, eg 75V, to obtain better resolution. The restriction site for the enzyme HaeIII has been given above. Predicted HaeIII fragment sizes for the ITS2 of different Gyrodactylus species are given in Table 6.


54

Restriction sites and fragment sizes can be predicted by analysing the DNA sequence by eye, or using suitable software.

TABLE 6 Gyrodactylus ITS2 ribosomal DNA HaeIII restriction fragment sizes.

Fragment sizes (bp) Enzyme

G. salaris G. teuchis G. derjavini G. truttae

556

HaeIII 321

234 234 234

167 167

154

63

46

45 Sequence analysis of the V4 and ITS ribosomal DNA regions While probe hybridisation and RFLP give some information about differences in the DNA of different species, they will not be able to detect all differences. Sequencing of the V4 and ITS PCR products will give the maximum amount of information as the complete DNA sequence is obtained. DNA sequencing is not suitable as a routine diagnostic method, principally due to the greater expense and time required. However, in the case of specimens with unusual morphological features, or aberrant hybridisation or RFLP patterns, it is useful to obtain the entire sequence as this will clearly indicate the level of differences and help define the relationship of the specimen to characterised species. ITS sequences have now been obtained for over 30 species of Gyrodactylus and have been useful for species identification and phylogenetic analysis (Cable et al., 1999; Cunningham et al., 2000; 2001; Matejusová et al., 2001; Zietara et al., 2000). Sequencing protocols can be found in many textbooks and manufacturer’s protocols. PCR amplified DNA can be sequenced directly or following cloning. It is preferable to obtain sequences from both strands of DNA and from replicate independent amplifications from each lysate, to ensure a reliable consensus sequence is obtained. Sequences can be compared to those deposited on public databases (EMBL, GenBank, DDBJ). In our experience, the degree of variation expected between Gyrodactylus species is of the degree found between the species shown in Figures 10 and 14 and summarised in Table 7 below. Very little intraspecific variation in V4 and ITS sequences has been found in Gyrodactylus species analysed to date by the authors. This data is summarised very briefly in Table 8. Differences in one or two nucleotides have not usually been considered sufficient to delineate different species. The one exception to this is the case of G. salaris and G. thymalli as discussed below (see case studies).


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TABLE 7 Pairwise comparison of small subunit ribosomal RNA gene V4 region and ribosomal RNA gene ITS1 and ITS2 from Gyrodactylus salaris, G. teuchis, G. derjavini, and G. truttae. The V4, ITS1 and ITS2 regions of G. thymalli are identical to those of G. salaris and are not included in the table. Indel: insertion or deletion.

Salaris/ Derjavini

Salaris/ Truttae

Salaris/ Teuchis

Derjavini/ Truttae

Derjavini/ Teuchis

Truttae/ Teuchis

V4

% divergence 3.3 9.3 3.3 10.0 6.7 6.0

Transitions 4 5 2 7 6 3

Transversions 1 9 3 8 4 6

Indels 0 0 0 0 0 0

ITS1

% divergence 19.0 15.8 22.4 20.1 26.2 22.1

Transitions 24 20 33 16 24 25


Indels 59 45 73 79 104 71

ITS2

% divergence 3.8 6.9 3.6 5.9 6.1 6.9

Transitions 5 17 6 13 12 16


Indels 0 0 0 1 0 1


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TABLE 8 Intraspecific variation in Gyrodactylus rDNA sequences.

G. salaris

Sequence Total no parasites Total no sites Total no

countries Maximum

variation (bp) V4 10 5 4 0

ITS1 17 5 4 1

ITS2 11 8 3 2

G. teuchis


countries Maximum


ITS1 12 5 4 0

ITS2 8 4 4 0

G. derjavini


countries Maximum


ITS1 96 31 5 1

ITS2 30 16 4 2

G. truttae


countries Maximum


ITS1 30 10 3 1

ITS2 12 2 2 0 Integration of morphological and molecular diagnostic methods The principal advantages and disadvantages of both diagnostic approaches, morphological and molecular, have been discussed in the Introduction. In the case of unknown or uncharacterised species, sole reliance on one approach or the other can prove misleading. A vast number of morphologically defined Gyrodactylus species still remain uncharacterised by molecular means. In addition, new species may still be undescribed. Identical probe hybridisation or RFLP patterns may be found in two or more species, even though the overall DNA sequence of the V4 or ITS may differ significantly, as was found with G. salaris and G. teuchis (Cunningham et al., 2001). Likewise, it is possible that different Gyrodactylus species could share near identical morphological features, as in the case of G. salaris and G. teuchis, making identification by this method extremely difficult. Therefore, it would be unwise to depend on one form of identification only when both together can either confirm identification or highlight differences.


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Outline method for integrated identification of Gyrodactylus parasites Firstly, as mentioned earlier, only live or ethanol preserved parasites should be used. Formalin fixed parasites make molecular analyses difficult. Prepare living or ethanol preserved parasites for morphological examination as described previously, with removal of the opisthaptor from the body.

• Place a drop of water on a microscope slide. • Remove parasite from fish with forceps or needle, wash or blot the parasite to

remove excess ethanol, and place in the drop of water. • Using a scalpel blade or other fine pointed instrument, detach the opisthaptor from

the body. • Transfer the body to a 0.5 ml microfuge tube for lysis as described previously, and

subsequent use in V4 probe hybridisation or ITS PCR. • Fix the opisthaptor with Malmberg’s fixative, under a coverslip, as described

previously. • Examine morphological characteristics. Alternatively: • After washing or blotting the live or ethanol preserved parasite, place it in a drop of

water on a microscope slide. • Place coverslip on top of parasite. • Soak up excess water by holding a piece of filter paper to the edge of the slide. • Examine the morphological characteristics, identify, and/or photograph the specimen

for later examination. • Do not allow the specimen to dry out while under examination. Add more water to

the specimen with a pasteur pipette placed at the edge of coverslip if necessary. • Remove the coverslip from the slide, all the while observing the procedure under a

dissecting microscope. This will allow you to see if parasite sticks to coverslip or slide surface.

• With a fine forceps remove the parasite and place in a 0.5 ml microfuge tube containing lysis buffer.

• Process for V4 probe hybridisation or ITS-RFLP as described previously. One advantage of the first method is that Malmberg’s fixative can be used to study the opisthaptor. This gives better visualisation of the morphological characters of hooks, anchors and ventral bar. A permanent mount of the specimen is also kept, which can be examined in more detail later if molecular results prove interesting. Morphological analysis does not have to be carried out immediately, if time is limited. It can however be difficult to separate the opisthaptor from the body, and one or both parts may be lost. Care must also be taken with the second method to avoid loss of the specimen. An overview of the methods used in identification of Gyrodactylus species, as discussed thus far is given in Figure 17.


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Figure 17: An overview of methods used in the identification of Gyrodactylus species.

Mount parasiteson slide

Digest tissues

Catch live fish

Transport alive Preserve in ETOHFix in formalin

Kill fishprior to examination

Dissect into sections

Examine fish under microscope

Examine morphological features

Lyse parasite

Hybridise probes

Compare patterns with controls

Sequence DNA

Chaetotaxy

V4 PCR ITS PCR

ITS RFLP

Remove parasites from fish


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Case Studies The case studies given below are examples of problematic species identification when molecular or morphological techniques were used alone. When both approaches were used together, characterisation was possible. G. salaris/ G. teuchis (see Cunningham et al., 2001) Specimens of a Gyrodactylus parasite infecting rainbow trout were analysed morphologically and found on first examination to strongly resemble G. salaris. V4 probe hybridisation was carried out and it too indicated that the parasites were G. salaris, ie the G. salaris V4 probe bound to the specimen DNA (Johnston et al., 1996). However, when the ITS region was amplified and digested using the restriction enzyme Sau3A, it gave the restriction pattern normally found for G. derjavini. If only one method had been used to identify the parasite, then it may have alternately been classified as G. salaris or G. derjavini. Due to the conflict in results arising from different diagnostic methods, the specimens were examined more closely. Morphological studies showed that, while it did closely resemble G. salaris, there were differences in some of the morphological characters. DNA sequencing of the V4 and ITS regions revealed sequence differences of the same order of magnitude as that found between other clearly defined Gyrodactylus species (Table 7). The V4 probe for G. salaris had bound to the specimen DNA because of sequence homology between the two species in the region of the probe, but other regions of the V4 differed significantly. This new species was named G. teuchis (Lautraite et al. 1999; Cunningham et al. 2001). It can be successfully separated from G. salaris by differences in its ITS RFLP pattern, using the restriction enzyme HaeIII. Even allowing for perfect slide preparations and clear species characteristics, morphological features do not always reflect changes at a sub species level, which may have important consequences for fish health and disease management, such as development of pathogenic strains within a species. RFLP analysis and DNA sequencing offer additional methods by which these changes might be observed. G. salaris variant Gyrodactylus parasites from S. trutta from a number of different rivers within Denmark were analysed. The ITS region was amplified and digested with the restriction enzyme HaeIII. It gave a restriction pattern distinct from those Gyrodactylus parasites for which the HaeIII restriction pattern has been characterised, ie G. salaris/G. thymalli, G. teuchis, G.derjavini and G. truttae (Buchmann et al., 2000). The ITS region was sequenced and it was found to be very similar to that of G. salaris, with just three nucleotide differences. Morphological analysis again showed very high similarity to G. salaris. It was considered that these parasites represented a G. salaris “strain”. Infection studies were carried out and the “strain” was found to be non-pathogenic to salmon populations previously shown to be highly susceptible to the classical form of G. salaris and to demonstrate clear preference for O. mykiss hosts (Lindenstrøm et al., 2000). Thus a combination of different approaches in characterising these parasites revealed possible differences in pathogenicity between different strains. This finding may have important implications for management of gyrodactylosis.


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G. salaris/G. thymalli (see Sterud et al., 2002) G. salaris has a wide host range and is extremely pathogenic to certain strains of Atlantic salmon (Salmo salar). G. thymalli is a parasite of grayling (Thymallus thymallus), and though it can infect and reproduce on Atlantic salmon, it is quickly eliminated from this host. G. salaris and G. thymalli are considered separate species based on their host preference and pathogenicity. They are however extremely difficult to separate morphologically. In addition, sequencing of the ITS and V4 regions of the ribosomal genes has shown these two areas of the genome to be identical in the two species. The intergenic spacer (IGS) region of the ribosomal RNA gene array, which separates the 28S large subunit gene from the 18S small subunit gene, has been sequenced from G. salaris and G. thymalli (Collins & Cunningham, 2000). This region was analysed because it is not transcribed, and as such may be under less pressure to conserve its sequence. A higher number of mutations may accumulate in this region, between different species, compared to genes and spacers that are transcribed and further processed by the cell. Overall, the IGS was very similar between G. salaris and G. thymalli. A region of 23 bp DNA repeats was found within the IGS of G. salaris and G. thymalli. These repeats are not exact copies of each other, but can differ from each other at one to six positions. This region showed intraspecific differences in number of repeats and in repeat sequence. It also showed consistent differences between most, but not all, G. salaris and G. thymalli parasites analysed (Sterud et al., 2002) and may contribute to the overall diagnosis. The question of whether G. salaris and G. thymalli are two separate species or subspecies, is a contentious one. At the present time, an accurate diagnosis between G. salaris and G. thymalli can only be made by someone with a high level of expertise in morphological identification of Gyrodactylus parasites, and with knowledge of the host fish species from which the parasites were collected. The authors consider that development of the most reliable and informative diagnostic criteria is best achieved by an integrated approach, combining morphological and molecular features, and, on a wider scale; host, host susceptibility, and any pathogenic effect of the parasites. Additional Methods for the Identification of Gyrodactylus Species There are a number of other methods that have been developed to aid the identification of Gyrodactylus species, based mainly on characterisation of morphological features. Release of opisthaptoral attachment structures from tissue As an alternative to looking at the parasite or opisthaptor as a whole, the tissues surrounding the hard parts of the opisthaptor can be digested away. Tissue can often obscure structures, or prevent adequate flattening of the diagnostic features, making the shape and size of the structures difficult to see and measure accurately. If a suitable digestion buffer is used, the digested tissue can be collected and used in molecular diagnostic techniques, thus allowing analysis of the parasites by both methods as recommended above. Harris et al. (1999) used a technique for releasing opisthaptoral hard parts of individual ethanol preserved parasites by digestion of the surrounding tissues. All steps were carried


61

out using a stereomicroscope to ensure that hooks or anchors were not lost. The procedure was as follows: • Cut the opisthaptor from the body and place in a drop of water on a disc of film, 5 mm

in diameter. The discs are cut from an overhead projector transparency film with a hole punch and held on a slide with a small drop of water.

• If alcohol preserved, allow the opisthaptor to rehydrate for 10 minutes in the drop of water.

• Remove the water with a finely drawn glass Pasteur pipette. • Add 25 µl of distilled water. • Add 2.5 µl of 10x digestion buffer (75 mM Tris-HCl, pH 8.0, 10 mM EDTA, 5%

sodium dodecyl sulfate (SDS) and proteinase K (to a final concentration of 100 µg/ml).

• Incubate at 50ºC for up to 10 minutes, intermittently observing progress using the stereomicroscope.

• Remove digestion buffer and replace with distilled water. • The digestion buffer can be used in molecular diagnostic techniques as described

above. • Place a coverslip over the preparation and mount in ammonium picrate glycerine. A method has also been developed by Mo and Appleby (1990), modified from Yazaki (1982), to free the hard parts of the Gyrodactylus opisthaptor from surrounding tissue. This method, described below, was developed using live parasites. • Kill infected fish by inserting needle into the brain via the upper part of the eye. • Leave for approximately two hours in a shallow container, covered in water. The

parasites will detach from the dead fish and sink to the bottom of the container. • Collect parasites from the bottom of the container. • Transfer to a pointed glass tube, suitable for centrifugation. • Add artificial gastric juice (0.7 ml concentrated HCl, 0.1 g pepsin, in 100 ml distilled

water). • Digest worms at 37ºC for 24 hrs. • Centrifuge the digest preparation at 6000 rpm to pellet the undigested hard parts of

the opisthaptors. The hard parts will sink by themselves but centrifugation saves time.

• Siphon off the digestive juice. • Rinse the sediment, containing the hard parts, five times in distilled water, spinning

down the hard parts between each rinse. • Resuspend the sediment in a small volume of water and transfer a drop to a round

coverslip (diameter 10 mm). • Examine under a light microscope. • If hard parts are present, allow mount to air dry. Mo and Appleby (1990) used the method below to digest single specimens of Discocotyle sagittata but single specimens of Gyrodactylus could similarly be digested, taking extra care due to their smaller size. • Cut the opisthaptor from the body of the parasite. • Place the opisthaptor on a coverslip on a microscope slide so that digestion can be

monitored under a light microscope. • Add a drop of digestive juice.


62

• Digest at room temperature, intermittently observing progress under the light microscope. Digestion is usually completed in less than an hour.

• If necessary, very fine needles can be used to free the hard parts from the last remnants of tissue.

• Rinse the hard parts gently, using a syringe, five times with distilled water. • Allow mounts to air dry. Shinn et al. (1993) used the above method to release the hard structures of the opisthaptor from Gyrodactylus parasites. They recommended keeping the amount of digestive juice used to a minimum, (3-5 ml) as undissolved particles in the juice itself may be concentrated during the centrifugation step and interfere with subsequent stages. They suggested that the digestive juice could be filtered before use to avoid this problem. The above authors used a 1 in 1,500 solution of 2-phenoxyethanol as an anaesthetic to aid dislodgement of the parasites from the fins. If parasites were picked directly from fish, any adhering fish mucus was washed off in 0.2 M phosphate buffer before digesting. Release of hard parts from alcohol fixed specimens using the above method required an incubation of 72 hours at 37ºC compared to 24 hours for live parasites. The preserved specimens were washed several times with distilled water before digestion. The above method was not successful when using formalin fixed parasites, even beyond the incubation times used for alcohol preserved specimens. A two step method was devised for formalin fixed specimens which resulted in the release of a large proportion of marginal hooks, although the anchors required further treatment to completely remove some remaining tissue. This method is given below: • Wash formalin fixed specimens for two hours in distilled water to remove fixative. • Digest for 24 hours at 37ºC in artificial gastric juice. • Centrifuge parasites at 6000 rpm for five minutes. • Decant supernatant. • Resuspend pellet in distilled water. • Centrifuge at 6000 rpm for five minutes. • Decant supernatant. • Resuspend pellet in 0.25 mg/ml trypsin (pancreatic trypsin) in 0.0016 M Tris pH 7.8

containing 0.049% BSA. • Digest at 37ºC for 24 hours. • Centrifuge parasites at 6000 rpm for five minutes. • Decant supernatant. • Resuspend pellet in distilled water. • Centrifuge at 6000 rpm for five minutes. • Decant supernatant. • Resuspend pellet in artificial gastric juice. • Digest at 37ºC for 24 hours. • Centrifuge parasites at 6000 rpm for five minutes. • Decant supernatant. • Resuspend pellet in distilled water. • Repeat previous three steps several times. • Agitate the final pellet. • Prepare mounts as previously.


63

Shinn et al (1993) have used sonication to release opisthaptoral hard parts from Gyrodactylus species. This was successful with fresh or frozen parasites but failed in the cases of alcohol preserved or formalin fixed parasites. The following method for sonication is taken from the above publication: Fresh or frozen parasites were placed in pointed glass centrifuge tubes in 3 ml of distilled water. The parasites were sonicated on ice using one of two methods: • Three bursts of 9 sec at a low peak to peak amplitude of 9.5 µm were used in a MSE

150 W ultrasonic disintegrator (point sonicator) at a nominal frequency of 20 Khz fitted with an exponential titanium probe. The resultant ultrasonic energy was concentrated by increasing the amplitude of vibration during the process by use of a conical probe (end diameter 1/8”) through which the ultrasound was emitted.

• The second method used was a sonic water bath (Kerry Pulsatron 125) connected to a 240 V power supply with a continuous power output of minimum 100 W. This was operated at a frequency of 40 KHz.

• Released opisthaptoral hard parts were collected by centrifugation, rinsed in distilled water and mounted on round coverslips as previously.

• Sonication time varied widely between samples. Times of 1-25 minutes were used.

The sonic water bath was preferred over the point disintegrator because the percentage of individual hooks retrieved using the water bath was 70-80% as opposed to just 10% and the resultant sonic energy was more controllable. Sonication also proved a better method when observation of musculature and ligaments associated with the opisthaptoral hard parts was desired. These remained in some sonicated specimens but were completely digested in the gastric juice treated specimens. Scanning Electron Microscopy (SEM) Scanning electron microscopy allows surfaces to be examined in minute detail. It has been used by a number of researchers in the study of the opisthaptoral hard parts of Gyrodactylus parasites (Mo and Appleby, 1990; Shinn et al , 1993; Harris et al., 1999). The opisthaptoral hard parts can be isolated by any of the above methods, after which the following steps, as given in Harris et al. (1999) are carried out to prepare the specimen for SEM. • Following digestion, air dry the preparation. • Examine the preparation using light microscopy to locate the anchors, marginal

hooks and bars. • Rinse gently and repeatedly with distilled water until all traces of buffer salts have

been removed. • Vacuum dry the specimen for 1-2 hours to ensure complete dryness. • Mount the film disc (or coverslip) on an aluminium stub using double sided carbon-

impregnated adhesive tape (Agar Aids, Stansted, UK), (or glue to stub with silver (Mo and Appleby, 1990)).

• Sputter coat with a gold palladium mixture. The specimen is now ready for analysis using scanning electron microscopy.


64

Statistical Classifiers This involves the application of statistical techniques to morphometric measurements obtained from parasite specimens via light and/or scanning electron microscopy. Based on one or a combination of measurements, parasites can be identified to species level or at least allocated to distinct groups. Good progress has been made in developing automated systems based on statistical classifiers. More information on this technique and its success with respect to identification of Gyrodactylus parasites can be found in Kay et al. (1999), McHugh et al. (2000), and Shinn et al. (2000). Chaetotaxy This is a technique that involves staining argentophilic surface sensory structures on the parasite with silver nitrate. When stained, these structures can be visualised easily under the microscope and differences in their arrangements and numbers can be used to differentiate between species. A discussion of this technique, including methodology can be found in Shinn et al. (1997; 1998a; 1998b). A brief overview of the methodology as outlined in Shinn et al. (1997) is given below: • Remove fins infected with gyrodactylids and wash briefly in 0.2 M phosphate buffer

pH 7.0 to remove excess mucus prior to staining. Alternatively, single parasites can be removed and washed but this increases risk of damage to the parasite surface.

• Parasites can be processed in situ on the fins. This means that all parasites are treated simultaneously, enabling standardisation of the silver nitrate treatment. Alternatively detached parasites can be processed individually.

• Place fin or parasite directly into 0.5% silver nitrate at 65-70ºC in the dark for five minutes.

• Wash fins/parasites in 5-10 changes of distilled water. • Submerge fin or parasite in distilled water and expose to UV light (325 nm) for five

minutes (each side in the case of fins). • Wash fins or parasites in several changes of distilled water. • Place fins or parasites in a solution of 90% alcohol and 10% glycerine. • Allow the alcohol to evaporate, leaving the fins or parasites in glycerine. • Select a parasite (pick the parasites off fins) and mount in glycerine on a glass slide. • Position the parasite dorso-ventrally and straight so as to maximise the number of

sensillae visible in any one focal plane. Use an excess of glycerine when mounting the parasites to allow enough depth to carefully roll the parasite in order to achieve the correct orientation and to prevent the parasites from being flattened too much under the weight of the coverslip.

• Draw a chaetotaxy map from the specimen with the aid of a drawing tube attached to the microscope.

• Compare with known patterns • Store slides in the dark at 4ºC Patterns are still discernible on slide preparations maintained in the dark at low temperatures after six months. In the case of Gyrodactylus parasites, it is believed the chaetotaxy patterns remain constant throughout life, unlike some other monogenean parasites. Chaetotaxy techniques are relatively simple and quick to perform. Chaetotaxy cannot distinguish all Gyrodactylus species (Shinn et al., 1998a). However, it has been used to separate G. salaris from the commonly found salmonid parasites, G. derjavini and G. truttae. The ability to distinguish G. salaris from G. teuchis has not been investigated. One other particular problem encountered with this technique when used with Gyrodactylus parasites,


65

is their viviparous nature. The presence of an embryo adds bulk to the specimen preventing complete flattening and easy discrimination of sensillae on the under surface.

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Bakke, T.A. and Jansen, P.A. 1991b. Susceptibility of Grayling (Thymallus thymallus) to

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Documents

DIAGNOSIS OF GYRODACTYLUS (MONOGENEA; FISH IN … · Other Gyrodactylus species commonly found on salmonids in Europe include Gyrodactylus derjavini Mikailov, 1975, Gyrodactylus truttae