Internal parasites continue to be a major cause of lost productivity and deaths in sheep and goats in higher rainfall areas of New South Wales, and occasionally in good seasons in pastoral areas (Lane et al 2015, Anon 2016).
Techniques for performing field total worm counts were developed in 1950’s (HMcL Gordon, pers comm) and later refined and published (Arundel 1967). Despite this, some field veterinarians make only a cursory examination of the gastrointestinal tract (GIT) during routine autopsy, and may be surprised when a laboratory reports high worm activity in samples submitted. In addition, diagnosis of rapidly acquired worm infections such as acute Nematodiriasis requires a thorough post-mortem examination, as worm egg counts at this stage are useless.
With practice, a field total worm count on a sheep or goat adds less than ten minutes to a routine autopsy, far less time than to collect and submit laboratory samples, and requires minimal equipment. It provides immediate valuable insight to the current worm status of the animal and implications for recent drenching and grazing management of the mob.
In addition to routine autopsy tools, additional equipment for a field total worm count comprises (Figure 1):
The reagent bottles fit inside the specimen jars, and these and the funnel and spare string fit inside the sieve jar for storage and transport.
Select a suitable candidate, typical of the problem being investigated. While a moribund or dead sheep or goat is often the only specimen a producer is willing to sacrifice, it may not be ideal.
Avoid contamination of the GIT with dirt and grass. Choose a green grassy site in preference to bare dirt or gravel. Some prefer the tray back of a ute. Lay out the equipment, placing the opened sieve jar on the white tray. Opening the carcass in right lateral recumbency gives immediate access to the abomasum, but use your standard autopsy approach.
Don’t faff about poking and pulling the gut, particularly the abomasum, which dislodges worms to adjoining areas.
Use warm water. It helps reduce viscosity of mucus, improving fluid flow through the sieve.
Samples of intestinal content for culture or isolation of epsilon toxin and of tissues for histopathology are best obtained before proceeding with the worm count.
Tie off the abomasum at both ends with string. (With practice, an alternative to string is to clamp the pylorus between the fingers of one hand to prevent escape of its contents while removing the abomasum.) Gently separate the abomasum from the rumen and tear the omentum. Elevate and loosely clamp the abomasum at its junction with the omasum with your free hand (if you haven’t tied it off) and separate with scissors. Remove the abomasum from the animal continuing to hold with your free hand to prevent escape of contents, and transversely cut the duodenum immediately distal to the ligature.
Suspend the abomasum, placing the pyloric end into the opened sieve jar, and incise the lower end of the greater curvature with scissors to release its contents. The volume of the abomasal content is usually less than one litre. Be aware that particularly when carrying a heavy burden of Haemonchus, the contents of the abomasum of adult sheep might exceed the one litre capacity of your sieve jar. If this happens, take an aliquot from the first full sieve jar, discard the remainder and continue as normal, remembering to account for the changed dilution factor.
Complete the incision of the greater curvature, lay the opened abomasum on the tray, and remove all worms from the mucosal folds with small volumes of water using the back of the hand as a scraper.
Make up the contents to the nearest mark on the sieve jar, invert the jar a couple of times to thoroughly mix and suspend the contents using one hand as a lid, and remove two 50ml aliquots. Discard the remainder.
Place one of the aliquots in the sieve jar (keep the second aliquot as a back-up), fill with warm water, secure the lid and sieve the contents. Add more water and repeat sieving until the contents is clear and about 50mls remain. Back-flush the lid with water to remove any worms clinging to the mesh, remove the lid, and place a few drops of iodine solution into the fluid.
Place this aside for a few minutes, and use the time to locate and free up the small intestine, tearing the duodenum and jejunum from the mesentery.
Return to the sieve jar, and decolourise the background only by adding hypo drop-by-drop. Try to be consistent in adding iodine and hypo. With experience, more faintly coloured worms are usually immature.
Spill the contents onto the white tray, to identify and count worms present. If the numbers of worms are excessive, return the entire contents of the tray to the sieve jar, make up to the nearest mark, and remove a 50ml aliquot. Spill this aliquot onto the white tray, and again identify and count the worms present.
Multiple by the dilution factor used to arrive at an estimated total abomasal worm count. For example, if the gross content of the opened abomasum after washing was 800ml, you added water to the 1000ml mark and took a 50ml aliquot to sieve, stain and count, the dilution factor is x20. If there were too many worms to count, you returned the entire sample on the tray to the sieve jar, added water to 500ml, removed and counted the worms in a 50ml aliquot, the dilution factor is x200.
The three worms found in the abomasum are Haemonchus, Ostertagia and Trichostrongylus axei. They can be easily differentiated with the naked eye using length and thickness (Arundel 1967). Haemonchus are large worms, males up to 15mm long, and females about 25mm long. The latter have characteristic barber’s pole effect, the white reproductive tract wound spirally around the blood-filled intestine. Applying iodine often destroys this effect. Ostertagia are slender brown worms about 12mm long and uniform thickness throughout their length (“eyelash”). T.axei are about 6 to 7mm long, and taper markedly. To differentiate immature forms, which may be confused based on size alone, Arundel (1967) notes these lose their iodine stain quickly.
Remove the first 3.5 to 5 metres of small intestine, being careful at all times not to lose sight of the proximal (pyloric) end of the duodenum. Twelve to fifteen “hand-over-hand” lengths is about right. Use the opened carcass or place the white tray to ensure the freed intestine does not contact the ground.
If you have trouble finding the duodenum, remember that the middle section is attached ventrally to the descending colon. Don’t let go of the end of the duodenum. Securely hold the pyloric end of the duodenum between fourth and little finger of one hand, and carefully place the remaining duodenum and jejunum in the sieve jar to avoid kinks and knots.
Seal the inside of the funnel with a thumb or one finger and scoop up warm water from the bucket. Place the nozzle of the funnel into the opened pyloric end of the duodenum, releasing a dribble of water to help separate and lubricate the collapsed walls if necessary.
Add about 100ml warm water through the funnel, remove the funnel, return the free end of the intestine to your finger grip and massage the water bolus distally, ensuring the distal end of the small intestine remains in the sieve jar. Repeat the flushing process. Note the number and size if any tapeworms present.
Open the intestine longitudinally with blunt-end scissors and examine the mucosal surface for abnormalities that may indicate parasites, including thickening, a ground glass appearance, haemorrhage, congestion or coccidial nodules.
Gently scrape the mucosa by pulling the opened intestine between the scissor handles held above the sieve jar (Webb, 1988 pers comm). Gordon (1988 pers comm) also recommends collecting mucosal smears at this point for later microscopic examination for immature worms.
Fill the jar with warm water to the nearest mark, invert to thoroughly suspend contents, remove two 50ml aliquots and discard the remainder. Return one aliquot to the sieve jar, fill with warm water and sieve the contents twice, leaving about 50mls residual fluid. Back-flush the sieve lid with warm water before unscrewing. Add iodine to the jar as before. Allow to stand while examining the large intestine. Then decolourise the background with hypo. Spill the stained worms onto the white tray and identify and count the worms present.
Small intestinal worms include Trichostrongylus, Nematodirus, Stongyoides and Cooperia. Trichostrongylus spp are small, slender, strongly tapering worms to about 7mm long. Nematodirus are much longer, females to about 23mm long and males to about 15mm. Their “thin neck” is usually coiled and males stain poorly. Strongyloides are small, poorly-staining and about 6mm long. Take care to differentiate from immature Trichostrongylus. Cooperia, reddish worms about 15mm long and often coiled like a watch-spring, and of uniform thickness, are rarely present in large numbers in sheep.
A thorough worm count of large intestinal worms is not attempted in the field. It requires large volumes of water and high-capacity sieves. These worms are rarely the only or the most pathogenic in the animal. They are large and the burden roughly assessed by opening the caecum and proximal colon.
Both Chabertia and Oespophagostomum are white-cream, about 20mm long. Chabertia have their trade-mark large mouth, while Oesophagostomum tapers at both ends (Arundel 1967).The 35-60mm whip-shaped Trichuris is usually found attached to the wall of the caecum. In our area, a large number of Trichuris worms is often associated with high stocking rates and little paddock feed; Trichuris L3 larvae remain in the egg shell, requiring sheep to eat the egg to continue the life cycle.
In deciding what number of worms is significant, Gordon (1981) observes that the pathogenic effects of worm species vary; that host factors such as age and body condition influence susceptibility to a given worm burden; that the duration of infection impacts the effect of a given worm burden; and that mixed species infections often have additive effect.
Total undifferentiated mixed worm infestations are arbitrarily divided into low, medium and high categories by Kingsbury (1965), using counts of <4000, 4000-10000, and >10000 worms respectively. While this system provides a baseline for beginners developing expertise with species identification, it tends to overlook the relative pathogenicity of the various species.
Recognising these differences, Gardiner and Craig (1961) assess pathogenic importance of mixed infections with a points system:
Ostertagia 3000 worms = 1 point
Haemonchus 500 worms = 1
Trichostrongylus axei 4000 worms = 1
Trichostrongulus spp 4000 worms = 1
Nematodirus 3000 worms = 1
Chabertia 100 worms = 1
Oesophagostomum 100 worms = 1
Immature worms (all spp) 4000 worms = 1
The worm burden is significant in any weaner sheep with a score greater than 2 points.
McKenna (1981) modified this scoring system, arriving at a “total pathogenic index”. While comprehensive, more calculations are needed.
The numbers of single worm species producing heavy infestations and mortalities in sheep have been assessed (Table 1, Eamens 1985).
Table 1. Worm species producing heavy infestations (mortalities)
|WORM SPECIES||YOUNG SHEEP||ADULT SHEEP|
|Trichostrongylus||5,000 (poor weaners)||30-40,000|
|20,000 (strong weaners)|
Estimating the number and type of gastrointestinal worms during a sheep or goat post-mortem examination is quick and enlightening. As with any skill, the technique requires practice which is best gained from making it part of every routine post-mortem.