20 for 20: Day 7 and day 8: Author in dauer state

Mata ne !  Come back soon.

20 for 20 : Day 6 : Calculating local HDUs (Heartworm Development Units)

While trying to define heartworm development units (HDUs) for veterinary students in the year two Veterinary Parasitology course, I chanced upon a 2015 article in Veterinary Parasitology by Ledesma and Harrington, titled “Fine scale temperature fluctuation and modulation of Dirofilarial larval development in Aedes aegypti”. The HDU is a calculated measure that factors in temperature to determine the extrinsic incubation period of heartworm larvae in mosquitoes.

Mosquitoes are obligate biological vectors of  Dirofilaria immitis. They bite infected dogs, and take up circulating microfilaria. These microfilaria have to develop to the third stage larval form (called L3) in the mosquito before they can infect dogs and mature to adulthood. The maturation from microfilaria to L3 takes time and is dependent on temperature. The warmer it is, the faster the larvae mature, with no maturation taking place below 14 degrees Celsius. This is expressed as the HDU, with at least 130 HDUs  being necessary before the L3 migrate back to the mosquito head and labium, ready to infect the next animal that the mosquito feeds on.

The formula for calculating the HDU is ∑ Average daily temp - 14°C = Accumulated HDUs.

As I was preparing for my lecture tomorrow, I thought it would be fun to calculate and compare the HDUs afforded by the climate in different seasons in the town that I live in. I obtained daily temperatures (in degree celsius, because life loves the SI system) from January (winter), April (spring) and August (summer) and made my calculations.

130 accumulated HDUs can be ruled out in January, and there is no development of microfilaria to the L3 stage in mosquitoes that reside outside. However, near and inside buildings and parking lots close to areas with centralized heating, in the so-called “urban islands”, the story might be different.

In march, if a mosquito picked up microfilaria on the first day of the month, infective L3s will have theoretically developed by day 14, when 130 accumulated HDUs are reached. Similarly, in august, if microfilaria had been picked up on the first, theoretically L3s will have developed by day 8.

In terms of actual transmission risk, real life in this case probably follows theory. There is lower risk of a dog contracting canine dirofilariasis in the winter, than in the spring, which is only slightly lower than summer in this town. It is the duty of every owner then to ensure that their dogs receive the chemoprophylactic prescribed by their vet at the right time and the right dose to prevent infection.

This post is part of a goal to write for 20 mins for 20 days.

20 for 20: Day 5 : From the “LAMP”-post to Cair Paravel on the Eastern sea

Once in a while, a diagnostic technique comes along that is everything you ever dreamed about, and wonderful like a Narnian summer. It is highly sensitive, highly specific, costs very little, does not require bulky equipment or highly trained personnel. One that has been hailed that way is the Loop mediated isothermal AMPlification assay or LAMP for short.

LAMP results in the amplification of nucleic acid sequences at a single, constant temperature (hence, isothermal). A special strand displacement polymerase, such as Bst polymerase from Bacillus stearothermophilus is used, along with 6 primers that confer specificity to the assay. Amplicons are later visualized using a fluorescent dye (increase in fluorescence indicates amplification), on an agarose gel or using a magnesium pyrophosphate (precipitation of the salt implies amplification). 

The LAMP assay at first glance seems to be all that it promises to be. It is specific and sensitive, costs less than conventional PCR, requires only a heat block or water bath at a constant temperature, and monkeys can probably be trained to do it. However, despite the hundreds of original research articles that have been published for parasite detection, no one seems to be routinely using it in diagnostics.

A little digging into the details of the assay brings the disadvantages to light.

(1) The LAMP assay is an end-point assay. One can measure the turbidity or fluorescence and try to correlate that to parasite number, but the amplicons themselves cannot be used for further cloning or sequencing, ruling out its use in the research lab.

(2) Designing primers for the LAMP may not be the easiest thing in the world, despite the availability of softwares. Since non-specificity at low annealing temperatures is always an issue when designing nucleic acid primers, designing 6 primers on a short sequence that all anneal at one temperature is pretty hard.

(3) Contaminants in the template inhibit amplification. If you have ever tried to amplify sequences from feces directly, you know what I am talking about. There are (probably) a million (I exaggerate) inhibitors in feces that impede conventional PCR even with a robust polymerase. These same inhibitors also impede the sensitive Bst polymerase as well. If you have to purify DNA using a kit, you might as well do conventional PCR/qPCR, and sequence the amplicons.

(4) Multiplexing is hard because of the numbers of primer pairs one will have to design. For example, duplexing the assay requires one to design 12 primers. A triplex assay requires 18 primers and so on.

The utility of the LAMP assay depends on what it is going to add to clinical diagnosis. For example, a veterinarian handling a sheep flock with trichostongylosis is probably going to help the farmer more by doing a FECRT than LAMP assays separately for Haemonchus, Cooperia, Ostertagia, Trichostrongylus, Nematodirus and any other common nematodes in the area.

Therefore, while the LAMP is a great assay to design, read about and think through, it’s practical value in veterinary parasitology may be pretty limited at present.

This post is part of a goal to write for 20 mins for 20 days.

20 for 20: Day 4: Diversity of parasites: hookworm edition

There is an astonishing diversity in the number of parasites that affect animals and humans. For example, there are at least 68 species of hookworms infecting 111 mammalian species, according to a new review article in the International Journal of Parasitology: Parasites and Wildlife titled “The diversity and impact of hookworm infections in wildlife”.

Hookworms are important blood sucking parasites of mammals, causing damage to the small intestinal mucosa to which they attach with their “hook”-like teeth/cutting plates. The parasites secrete anticoagulant proteins that cause the mucosal wounds to bleed, resulting in blood loss and hence anemia in the host, tissue damage, inflammation, retarded growth and even death. The larval stage, which is the infective form, can penetrate the host (animal and human) skin, and later migrates to its preferred site in the intestine.  

The paper systematically reviews the hookworms found in each mammalian order and family and presents the data in thirteen well-organized tables. You dear reader must refer to the original article for details (click here for the article). Here is the 10,000 feet view of the reported parasite genera affecting each mammalian family studied.

Canidae: Ancylostoma (A. caninum, A. tubaeformae. A. kusimaense, A. miyazakiense, A. buckleyi, A. ceylanicum, A. braziliense), Uncinaria (U. stenocephala, U. carinii)

Felidae : Ancylostoma (A. caninum, A. braziliense, A. tubaeformae, A. pluridentatum, A. buckleyi, A. paraduodenale), Uncinaria (U. maya, U. stenocephala, U. felidis), Galoncus (G. periniciosus, G. tridentatus), Arthrostoma (A. hunanensis)

Otariidae (eared seals): Uncinaria (U. hamiltoni, U. sanguinis, U. lyonsi, U. lucasi)

Procyonidae: Necator (N. urichi), Uncinaria (U. maxillaria, U. bidens), Arthorcephalus (A. lotoris), Ancylostoma(A. kusimaense), Arthrostoma (A. miyazakiense)

Mustelidae : Uncinaria (U. criniformis), Teteragomphius (T. procyonis, T. arctonycis, T. melis),  Ancylostoma spp.

Ursidae: Ancylostoma (A. caninum, A. tubaeforme, A. malayanum), Arthrocephalus (A. lotoris), Uncinaria (U. rauschi, U. yukonensis)

Mephtidae (skunks), Herpestidae (mongoose), Phocidae (seals), Hyenidae and Viverridae (civets): Ancylostoma (A. duodenale, A. ceylanicum), Arthrostoma (A. vampira, A. conepati), Arthrocephalus (A. lotoris, A. gambiensi), Uncinaria spp.

Bovidae : Agristomum (A. gorgonis, A. cursoni, A. monnigi, A. equidentatus), Bunostomum (B. phlebotomum, B. trigonocephalum), Gaigeria (G. pachyscelis)

Suidae and Tayassuidae : Globocephalus (G. urosubulatus, G. samoensis, G. longimucronatus, G. versteri)

Cervidae and Giraffidae: Bunostomum (B. phlebotomum, B. trigonocephalum), Monodontus (M. lousianensis), Monodontella (M. giraffae)

Primates : Ancylostoma spp., Necator (N. americanum, N. gorillae), Bunostomum spp.

Rodentia: Uncinaria (U. hydromyidis), Cyclodontostomum (C. purvisi), Acheilostoma (A. simpsoni, A. moucheti), Monodontus (M. floridanus, M. aguaiari, M. rarus)

Perissodactla, Proboscidae, Pholidota, Afroscoricida, Scandentia: Monodontus (M. nefastus), Brachyclonus (B. indicus), Grammocephalus (G. intermedius, G. clathratus, G. hybridatus, G. vardatus), Bunostomum (B. brevispiculum, B. hamatum), Bathmostomum (B. sangeri), Necator (N. americanus), Uncinaria (U. bauchoti, U. olseni)

Now, it is my duty to inform you that many of the above can affect humans. No worm is zoonotic till it is found for the first time in a human. Remember that the next time you go padding around barefoot on a distant beach or mountain.

Reference:
Seguel, Mauricio, and Nicole Gottdenker. "The diversity and impact of hookworm infections in wildlife." International Journal for Parasitology: Parasites and Wildlife (2017).

This post is part of a goal to write for 20 mins for 20 days.

20 for 20: Day 3 : Identity crisis with feline Trichuris

 At the WAAVP conference held in Kuala Lumpur this year, one of the last talks on the last day was delivered by Dr. David Elsemore from IDEXX. IDEXX has developed a battery of fecal antigen ELISAs to detect nematode infections in small animals. This particular talk was about the detection of cat whipworms by the antigen ELISA originally designed to detect the canine whipworm. One of the advantages of the whipworm antigen ELISA is that it detects excretory-secretory antigens produced by the live worm, but not egg antigen. So, any spurious parasites are not detected, eliminating false positives. The ES whipworm antigen used is apparently involved with maintenance of the infection in the host.

The talk got me thinking about Trichuris of felines, a rather uncommon parasite in the United States. A quick search on PubMed to look for recent articles brought up a SpringerPlus article by Dr. Jennifer Ketzis, titled "Trichuris spp. infecting domestic cats on St. Kitts: identification based on size or vulvar structure?". Dr. Ketzis covers the history of the taxonomy of the two species of feline Trichuris – T. serrata and T. campanula first. It turns out that there is a debate about the existence of two species, since early descriptions were based on size of the parasite and size of the eggs with some discrepancies occurring due to the examination of only a handful of worms in some cases.  After examining 96 male and 113 females, the researchers concluded that the Trichuris found on St. Kitts is T. serrata.

So, how does one distinguish between T. serrata and T. campanula?
a. Egg size is not definitive. Both speces have typically trichuroid eggs that are light brown, with bipolar plugs and a morula, but egg sizes overlap. Also, eggs recovered from feces and from adult worms vary in size.

b. Adult worm size is not definitive either, although that had been the defining feature in the original descriptions. Worm size according to Dr. Ketzis and others seems to depend on number of worms sharing the same host, host immunity affecting worm size in repeat infections, exposure to lots of infective eggs all at once resulting in smaller adults, and natural biological variation.

c. The only distinction between the two species is the presence of a finger like vulvar projection in the females of Trichuris serrata, but not Trichuris campanula, and the presence of a bacillary band in the former.

Although there are claims that the presence of many Trichuris in cats cause gross and histopathological lesions, Dr. Bowman in Georgi’s Parasitology for Veterinarians says that they are of “little practical importance” except to complicate the differential diagnosis of feline capillariasis.

Therefore, it is incumbent upon veterinary parasitologists to interpret with caution the presence of trichuroid eggs in feline feces, given that (a) they could be other capillarids, (b) there is no molecular data available to distinguish between T. serrata and T. campanula and (c) the antigen ELISA cannot distinguish between them either.

References:

Ketzis, Jennifer K. "Trichuris spp. infecting domestic cats on St. Kitts: identification based on size or vulvar structure?." SpringerPlus 4.1 (2015): 115.

Bowman, Dwight D. Georgis' Parasitology for Veterinarians. Elsevier Health Sciences, 2014.

This post is part of a goal to write for 20 mins for 20 days.

20 for 20: Day 2: The deal with Joyeuxiella

While flipping through the Bayer manual of  Helminthology in Veterinary Practice by Pachnicke et al., which has some very pretty parasite pictures, I came across the cestode Joyeuxiella. The name has connotations of a happy parasite, but was named after the French  veterinary parasitologist, Dr. Joyeux.

As I rooted around the internet and the books to learn about this parasite, I saw that besides being infected by the Cylophylideans, Taeniids, Mesocestoides and Dipylidium caninum, cats in other parts of the world except the Americas, can also be infected by two other genera in the Family Dipylidiidae, viz. Diplopylidium and Joyeuxiella. The latter two are very similar to Dipylidium in shape and size, except for the placement of genital apertures and egg morphology. Dr. Bowman’s book says that while the genital aperture is slightly behind the middle of the segment in Dipylidium, it lies anterior to the middle of the segment in Diplopylidium and Joyeuxiella. Also, while 5- 30 eggs are commonly observed in egg packets of Dipylidium, only one egg per egg packet occurs with Joyeuxiella and Diplopylidium.

Joyeuxiella uses coprophagous bettles as the first intermediate host, and reptiles and small mammals as second intermediate hosts. Cats and dogs can be definitive hosts. The genus was revised in 1983 (1) , and shrunk from 13 species to 3 valid species namely J. pasqualei, J. fuhrmanni and J. echinorhyncoides. The paper also presents the key to species identification in the genus.

Literature about Joyeuxiella is scant, and a search on PubMed brought up only 28 articles, of which 4 had been published between 2012 and 2017, and 14 between 2007 and 2017. An interesting older article published in 2006 was a feline case reported from Thessaloniki in Greece of intestinal pleating associated with J. pasqualei infection (2). The tapeworm had attached itself to the anterior parts of the small intestine, explaining the clinical signs associated with a linear foreign body. Geographically, incidence seems to cluster around the middle east, parts of Europe and Australia. In a study from Dubai (3), rather a large percentage (65.8%) of the cats trapped and euthanized seemed to be infected with Joyeuxiella species. In another study from Spain, feral cats were considered reservoirs of infection to Iberian lynxs (4).

Although the parasite seems absent in North and South America, it behooves Veterinary Parasitologists to be on the lookout, lest the parasite becomes established here, in yet another case of parasites taking to the skies and arriving in locations where they were not initially a problem.

References:
1 Jones, Arlene. "A revision of the cestode genus Joyeuxiella Fuhrmann, 1935 (Dilepididae: Dipylidiinae)." Systematic Parasitology 5.3 (1983): 203-213.
2 Papazoglou, L. G., et al. "Intestinal pleating associated with Joyeuxiella pasqualei infection in a cat." Veterinary record 159.19 (2006): 634.
3Schuster, Rolf K., et al. "The parasite fauna of stray domestic cats (Felis catus) in Dubai, United Arab Emirates." Parasitology research 105.1 (2009): 125.
4 Bowman, Dwight D. Georgis' Parasitology for Veterinarians. Elsevier Health Sciences, 2014.

This post is part of a goal to write for 20 mins for 20 days.