Susceptibility of Midge and Mosquito Vectors to SARS-CoV-2

Severe acute respiratory syndrome coronavirus 2, also known as SARS-CoV-2, is the notorious coronavirus responsible for the COVID-19 pandemic. As is the case with any disease, understanding how SARS-CoV-2 spreads through a population is one of the most important steps in slowing transmission. Early on, scientists determined that the primary mode of infection for SARS-CoV-2 is through exposure to respiratory droplets carrying infectious virus (CDC 2021). This means the most likely route of infection would be through direct, indirect, or close contact (<6 feet) of an infected person through infected secretions.

In 2020, scientists published findings that SARS-CoV-2 viral RNA (genetic material of the virus) was identified in the blood and serum of infected patients, suggesting that the virus could also be present in the bloodstream (Chen et al. 2020, Hogan et al. 2020, Young et al. 2020). This led researchers to question if blood-sucking arthropods like biting flies could also serve as a mode of biological transmission for SARS-CoV-2. Initial experiments testing this hypothesis confirmed that the virus did not replicate in three common mosquito species (Aedes aegypti, Ae. albopictus, and Culex quinquefasciatus) after the virus was injected directly into the mosquitoes (Huang et al. 2020). Additional research by Xai et al. (2020) showed that the virus did not replicate in cells collected from Aedes mosquitoes, and SARS-CoV-2 was never recovered from field-caught Culex or Aedes mosquitoes from Wuhan. These results were promising, but only a few species were tested, and researchers injected the mosquitoes with the virus rather than allowing them to ingest an infected bloodmeal. 

Balaraman et al. (2021) set out to address some of the holes left in previous research by testing if more species of biting flies were susceptible to the SARS-CoV-2 virus after ingesting an infected blood meal. The research team expanded on the list of biting insects investigated to include two previously untested flies, a biting midge (Culicoides sonorensis) and another species of Culex mosquito (Cx. Tarsalis), along with the previously tested Culex species Cx. quinquefasciatus.

To get a better idea of susceptibility of these insects to the virus, the research team measured the infection potential of these biting insects in two different ways. First, cultured cells from all three species were exposed to SARS-CoV-2 to determine if the cells from any species were vulnerable to infection or damage. Results from these experiments were negative for all insects, meaning they showed no signs of cellular infection or damage.

The next step was to test if any of the three species would be susceptible to infection after ingesting an infected bloodmeal. Researchers fed the insects on blood spiked with SARS-CoV-2 and held them for 10 days. After the waiting period, all blood-fed insects were tested for the presence of SARS-CoV-2 viral RNA and positive samples were set aside for further evaluation. It is important to note here that viral RNA is not the same as infectious virus. This means that simply confirming the presence of viral RNA would not also guarantee that infectious virus particles were present. To determine if any of the samples with viral RNA also contained infectious virus, another set of experiments were conducted that measured cellular damage to confirm viral infection. Those results were all negative meaning that none of the samples tested positive for infectious virus. 

It may seem like a no-brainer that the mosquito, an insect crowned the world’s deadliest organism because of the diseases it is known to spread, should also be able to biologically transmit the SARS-CoV-2 virus. But the disease transmission process can be extremely complicated. The overarching conclusion we can draw from these studies is that the biting insects studied so far most likely do not play a role in biological transmission of SARS-CoV-2. While we cannot rule out the possibility of arthropod-borne transmission of SARS-CoV-2 all together, there is no evidence so far to suggest this route of transmission is possible.

Research is currently ongoing to measure the potential for some insects (mostly cockroaches and flies) to mechanically spread the SARS-CoV-2 virus after contacting a contaminated surface or material. While this route of viral spread may be more plausible, the likelihood that a person contracts COVID-19 after contacting a contaminated surface is low CDC (2021).

Mike Bentley, PhD, BCE


References:

Centers for Disease Control and Prevention. (2020). Science Brief: SARS-CoV-2 and Potential Airborne Transmission. https://www.cdc.gov/coronavirus/2019-ncov/science/science-briefs/scientific-brief-sars-cov-2.html.

Centers for Disease Control and Prevention. (2021). Science Brief: SARS-CoV-2 and Surface (Fomite) Transmission for Indoor Community Environments. https://www.cdc.gov/coronavirus/2019-ncov/more/science-and-research/surface-transmission.html.

Chen, W., Y.  Lan, X.  Yuan, X.  Deng, Y.  Li, X.  Cai, L.  Li, R.  He, Y.  Tan, X.  Deng, . et al.  2020. Detectable 2019-nCoV viral RNA in blood is a strong indicator for the further clinical severity. Emerg. Microbes Infect. 9: 469–473.

Huang, Y. J. S., D.L.  Vanlandingham, A.N.  Bilyeu, H.M.  Sharp, S.M.  Hettenbach, and S.  Higgs. . 2020. SARS-CoV-2 failure to infect or replicate in mosquitoes: an extreme challenge. Sci. Rep. 10: 1–4. 

Hogan, C. A., B. A.  Stevens, M. K.  Sahoo, C.  Huang, N.  Garamani, S.  Gombar, F.  Yamamoto, K.  Murugesan, J.  Kurzer, J.  Zehnder, . et al.  2020. High frequency of SARS-CoV-2 RNAemia and association with severe disease. Clin. Infect. Dis. 1–5. 

World Health Organization. (2021). Malaria. https://www.who.int/news-room/fact-sheets/detail/malaria.

Xia, H., E.  Atoni, L.  Zhao, N.  Ren, D.  Huang, R.  Pei, Z.  Chen, J.  Xiong, R.  Nyaruaba, S.  Xiao, . et al.  2020. SARS-CoV-2 does not replicate in Aedes mosquito cells nor present in field-caught mosquitoes from Wuhan. Virol. Sin. 35: 355–358. 

Young, B. E., S. W. X.  Ong, S.  Kalimuddin, J. G.  Low, S. Y.  Tan, J.  Loh, O. T.  Ng, K.  Marimuthu, L. W.  Ang, T. M.  Mak, . et al.; Singapore 2019 Novel Coronavirus Outbreak Research Team. 2020. Epidemiologic features and clinical course of patients infected with SARS-CoV-2 in Singapore. JAMA  323: 1488–1494.

Finding Ways to Reduce the Risk of Non-Target Rodenticide Exposure in Roof Rat Control

Earlier this month, a research paper was published in the journal Pest Management Science titled “Use of Rodenticide Bait Stations by Commensal Rodents at the Urban-Wildland Interface: Insights for Management to Reduce Non-Target Exposure”. This research was partially funded by a Pest Management Foundation grant made to Dr. Niamh Quinn’s research program at University of California Agriculture and Natural Resources, South Coast Research and Extension Center in Irvine California. The Pest Management Foundation is an independent non-profit charity that provides scholarships for outstanding urban entomology students and funds structural pest control research at universities across the United States. All of the Foundation’s work is made possible by donations from pest control companies and individuals interested in advancing the science of structural pest control.

In recent years, anti-pesticide groups have placed pressure on pest management professionals regarding non-target exposures of wildlife to anticoagulant rodenticides – especially in California mountain lions, birds of prey in New England, and most recently in bobcat populations on Kiawah Island, South Carolina. The knee jerk reaction by many of these activist groups has been to ban the use of these products based on residue data that may show sub-lethal amounts of anticoagulants in non-targets. In California, a measure was successfully passed in December 2020 to ban most SGAR uses across the state, even by licensed professionals. Unfortunately, outright bans like these don’t take into consideration the important beneficial role that rodenticides play in managing these important public health pests and that instead of banning the products altogether, it’s important to first understand how non-targets are becoming exposed, then try to limit those exposures through common sense risk mitigation.

Although it is not known exactly how these non-target exposures are occurring, it is thought that native rodents like deer mice, wood rats, kangaroo rats and ground squirrels are likely entering stations and consuming rodenticide bait, then raptors, coyotes, pumas or other non-target predators are eating the non-target, non-pest, native rodents.

To better understand which native rodents are entering bait stations, Dr. Quinn and her colleagues, positioned two bait stations in more than 90 backyards in southern California.  One station was placed at ground level and one was positioned 3-5 feet off the ground.  Non-toxic commercial bait was placed inside each station, then digital cameras were focused on the two bait stations and every animal that interacted or entered the bait station was photographed.  More than half a million photos were captured during the study and here’s what was learned: 

  • Roof rats were present at more than 80% of the sites. House mice and Norway rats were observed much less commonly.
  • Native rodents (deer mice, wood rats, kangaroo rats and ground squirrels) were relatively rare, visiting or entering bait stations at only 13% of the sites. 
  • Native rodents were five times less likely to enter the stations that were positioned off the ground.
  • Roof rats, on the other hand, were equally likely to find, enter and interact with bait stations when positioned off the ground.

The authors made four specific pest management recommendations based on this research:

  1. PMPs should monitor bait consumption closely during the first few weeks to ensure that adequate bait is available to control populations.  High populations of rats can deplete baits quickly.
  2. On average it took roof rats 7-8 days to enter a station.  SGAR baits typically take 3-5 days to work, so control effects may not be seen for as long as two weeks after application.  This should be communicated to clients so that expectations can be managed.
  3. In areas where predator species like coyotes are common (that is, homes closer to natural areas and parks) care should be taken to clean up any rat carcasses that are found in the open.
  4. When roof rats are the target pest, positioning bait stations 3-5 feet off the ground can limit non target entry into the stations. 

Jim Fredericks, PhD, BCE

To read the full research paper visit: https://onlinelibrary.wiley.com/doi/abs/10.1002/ps.6345

To learn more about the other research projects taking place in Dr. Quinn’s lab visit http://ceorange.ucanr.edu/humanwildlifeinteractions/

To learn more about the kinds of research that the Pest Management Foundation funds, or to make a donation, visit www.NPMAFoundation.org .

Dung Beetles as Vertebrate Samplers

A biodiversity survey is an important tool that scientists use to gain a baseline understanding of organisms in an ecosystem. These surveys record the abundance (how many) and/or diversity (variety of animals) of species in the environment so scientists can monitor any change over time. The data collected from these surveys are the cornerstone to conservation research because collecting this data tells us which habitats and animals are being negatively affected and require protection efforts.

One challenge with biodiversity surveys is that the traditional sampling methods used to collect data are largely dependent on visual counts that are inherently biased. For example, a survey wanting to track the number of birds in a habitat may be limited by the researcher’s ability to visually spot and correctly identify that particular species. This is especially challenging when the target organism is rare or difficult to spot.

To address this challenge, scientists developed a new technique for large-scale biodiversity monitoring known as environmental DNA surveillance (Figure 1). Environmental DNA (or eDNA) is genetic material collected directly from environmental samples such as soil or water. Using this technique, scientists can identify the DNA of several organisms that have contacted the sample, offering a less-biased surveillance tool that is ideal for counting elusive or rare organisms. While this technique has its obvious advantages over traditional methods, one drawback is that water tends to be better than soil at preserving DNA which means aquatic environments are easier to survey than terrestrial habitats.

Figure 1. The overall workflow for environmental DNA (eDNA) studies with examples of organisms that have been identified from environmental samples. Image Credit: https://doi.org/10.1016/j.biocon.2014.11.019

Previous studies have investigated the use of blood-feeding invertebrates such as flies, leaches, and mosquitoes as alternative sources of eDNA, referred to as invertebrate-derived DNA (iDNA), to survey land-based animals (Beng et al. 2016, Robson et al. 2016, Nguyen et al. 2020). However, these organisms are often difficult to collect, limiting their use in large-scale surveillance programs.

Dung beetles are a widely distributed group of detritivores that feed on the fecal matter of terrestrial animals. Earlier experiments showed that certain cells found in mammal dung and ingested by dung beetles could be used as a source of identifiable DNA (Gómez & Kolokotronis 2016, Kerley et al. 2018). Since dung beetles are easy to collect in large numbers and are widely distributed, and a method for iDNA detection was already identified, this gave dung beetles an advantage over other invertebrates for their potential use in iDNA surveillance.

To further investigate the use of dung beetles as iDNA samplers, Drinkwater et al. (2021) set out to address two questions. How long could mammal DNA remain viable for identification in the gut of one dung beetle species (Catharsius renaudpauliani), and would it be possible to identify the DNA of multiple mammal species from the gut contents of several dung beetles? To address the first question, researchers fed 60 C. renaudpauliani on cow dung, then analyzed the gut contents of selected individuals at set time intervals ranging from 0 to 56 hours. They found that sufficient DNA could be recovered up to 4 hours after feeding, but the amount of DNA recovered dropped to zero at 9 hours post feeding. This meant that mammal DNA could be preserved long enough in wild dung beetles to be identified. And, the animal DNA that was identified from these dung beetles would have been from dung consumed within a short period of time before the beetles were caught. To address the second question, Drinkwater et al. (2021) trapped dung beetles in the field and evaluated their gut contents for mammalian DNA. They were able to confirm DNA of six different mammalian taxa, with three being identified down to species.

Collectively, these findings showed that dung beetles could serve as an easy to collect and widely distributed source of iDNA. This study takes us one step closer to better understanding a new source of invertebrate DNA in the quest for improved sampling methods. It’s the steppingstones of scientific discovery like those published by Drinkwater et al. (2021) that pave the way to developing more advanced surveillance tools that could one day change the world.      

In case you missed it, this publication was also briefly highlighted on this episode of the NPMA BugBytes podcast.

By: Michael Bentley, PhD, BCE

References:

Beng, K. C., K. W. Tomlinson, X. H. Shen, Y. Surget-Groba, A. C. Hughes, R. T. Corlett, And J. W. F. Slik. 2016. The utility of DNA metabarcoding for studying the response of arthropod diversity and composition to land-use change in the tropics. Sci. Rep. 6: 1–13.

Drinkwater, R., Clare, E. L., Chung, A. Y. C., Rossiter S. J., Slade, E. M. 2021. Dung beetles as vertebrate samplers – a test of high throughput analysis of dung beetle iDNA. BioRxiv 2021.02.10.430568.

Robson, H. L. A., T. H. Noble, R. J. Saunders, S. K. A. Robson, D. W. Burrows, And D. R. Jerry. 2016. Fine-tuning for the tropics: application of eDNA technology for invasive fish detection in tropical freshwater ecosystems. Mol. Ecol. Resour. 16: 922– 478 932.

Nguyen, B. N., E. W. Shen, J. Seemann, A. M. S. Correa, J. L. O’donnell, A. H. Altieri, N. Knowlton, K. A. Crandall, S. P. Egan, W. O. Mcmillan, And M. Leray. 2020. Environmental DNA survey captures patterns of fish and invertebrate diversity across a tropical seascape. Sci. Rep. 10: 1–14.

Take a Quiz to see if you Fear Spiders Like Some Entomologists!

Award ceremonies are typically not extremely exciting events. This is not to say that I don’t love them and enjoy fraternizing with my fellow industry folks in jubilant celebration. These events are entirely necessary for industries or organizations to recognize their most distinguished, brightest, or exemplary people. However, the Ig Nobel Prize awards held at Harvard annually falls into an entirely different bucket of award ceremonies that is both fascinatingly weird and delightful, and how I came about to learn that there are entomologists, like myself, that fear spiders. Dr. Richard Vetter won the Entomology prize for his paper “Arachnophobic Entomologists: Why Two Legs Makes All the Difference”.

Forty-one entomologists were selected to complete a “Fear of Spiders Questionnaire (FSQ)”, which is a standardized test used in psychology (Szymanski and O’Donahue 1995); and also rate their fear and disgust of spiders; and answer other questions regarding negative spider experiences and reasons of spider fear. The average score for these entomologists was a 28.2, so they were just somewhat adverse to spiders.

However, it gets interesting when you dive into the entomologists that scored high on the spider fear scale and would be considered arachnophobic. In regards to the statement “Spiders are one of my worst fears”, five entomologists gave the highest “totally agree” score.

A forensic entomologist who routinely handles maggots, gave spiders a maximum disgust score and replied “I would rather pick up a handful of maggots than have to get close enough to a spider to kill it.”

Nineteen of the 41 entomologists stated they had negative incidents with spiders. These happened both in youth and in adulthood. Respondents listed exposure to spiders crawling on them, bites or presumed bites, seeing large orb weavers in webs or running into them, having nightmares about them, and being tormented by family members or peers. Three entomologists specifically mentioned exposure to black widow spiders as one of their negative experiences.

One of the reasons entomologists described spiders negatively was because of their many legs – all spiders have 8 legs. I would laugh, if I were not a spider fearing entomologist, because we entomologists routinely handle 6 legged creatures. It seems those two extra legs, as the author’s title states “makes all the difference”. Also the real, or mostly perceived, notion of spiders running fast, showing up unexpectedly, having dangerous bites or just being ugly were all reported as why they evoke such negative emotions.  

If you are a pest management professional or entomologist that fears spiders – you are not alone! Arachnophobia usually develops in childhood. Most people don’t know the exact moment they became fearful but they have always been afraid of spiders. Entomologists who fear spiders have similar fears as the general public and not surprisingly, developed these fears way before considering entomology a career. So, I’ll continue to slay bed bugs all day but squeal like a child when I run into a spider web in a cold, dark crawl space.

Curious to see how you score on the Fear of Spider Questionnaire? Check this link to take the quiz. * We are bug doctors, not psychologists, so we can’t help you with any of your spider fear. We can only commiserate with you.

Brittany Campbell, PhD, BCE

Vetter, R. S. (2013). Arachnophobic entomologists: when two more legs makes a big difference. American Entomologist59(3), 168-175.

Szymanksi, J. and O’Donohue, W. (1995). Fear of spiders Questionnaire. Journal of Therapy and Experimental Psychiatry, 26(1), 31-34.

Undertakers and Corpse Disposal In The Termite Colony

In North America, termites cause more than $5 million in damage each year. Eastern Subterranean termites are the most widely distributed and most common encountered termite pest in North America. With colonies that can grow to the size of 5 million individuals and assuming a death rate similar to comparable termite species, it is estimated that as many as 70,000 termites could die in colonies each day. Corpse removal, a duty performed by worker termites called “undertakers” is extremely important for colony health. The longer a termite corpse remains, the greater the chances for disease spread. So, undertaker behavior (removal and disposal of dead termites), performed by worker termites,  is extremely important for colony health.

Eastern Subterranean termites have developed two distinct behaviors to deal with dead nestmates: cannibalism and burying the dead. Cannibalism or eating the dead, provides an important service to the colony by recycling nutrients back in the colony. Wood, which is the primary food for Eastern subterranean termites is notoriously lacking in nitrogen. By eating the dead, nitrogen-rich material from the corpses  is recycled back into the colony.  Additionally, gut symbionts, necessary for digesting cellulose may also be recycled. Burying behavior, on the other hand, ensures that the threat of entomopathogens and subsequent disease is eliminated from the colony, which is especially important for eusocial insects living inside the closed system. 

When a termite dies is begins to emit certain chemicals as it decomposes, including airborne volatile compounds such as 3-octanol and 3-octanone are released.  This tells the workers that they are dead and they need to be taken care of.  These chemicals do not persist for long and dissipate over time. Fatty acids are also produced, which are persistent. These chemicals build up in the corpse over time. The relative amounts of these chemicals in a termite corpse can provide information to undertakers to help determine what to do with the dead termite. New corpses were eaten if they were less than 64 hours old (higher levels of 3-octanol, 3-octanone) lower levels of specific fatty acids.

As a part of a series of laboratory experiments Jizhe Shi and his colleagues at the University of Kentucky learned the following information. Based on the levels of these chemicals encountered by undertaker termites (lower levels of 3-octanol and 3-octanone, and higher concentrations of fatty acids) older corpses were dealt with in two ways: either burying or walling off. All “old” worker corpses were buried or covered up with sand particles, feces, or other materials soon after discovery.

Interestingly, soldier termites were buried 50% and walled off about 50% of the time. When walled off, the entire tunnel to the chamber in which the dead soldier termites were present was blocked off.  This behavior seems to be adapted for situations in which the colony is breeched by invading ants/termites and large numbers of dead soldiers remain. Workers instinctively block off that entire area because it’s perceived as unsafe, and the dead termites present there serve as  reminder of danger if the area is opened up in the future.

It is notable that workers produce greater amounts of 3-octanol and 3-octanone upon death and were dealt with more quickly than soldiers. The authors of the study hypothesized that this was because of the greater proportion of workers present in the colony compared to other castes.  Due to sheer the numbers present, workers need to be cleaned up first. 

So, what does this mean for termite control? At first glance, the practical implications for our industry might not seem obvious – because they aren’t. This research does tell us that termites are not using visual cues like we might to determine the age of a corpse.  Remember when the gang from the Goonies found One-Eyed-Willie on his ship while being chased by the Fratellis?  They knew One-Eyed Willie was dead by looking at him, not sniffing him. Termites would have used chemical cues instead to know that he needed to be buried and not eaten.

This is the kind of research that might be categorized in the “cool to know” category, but it could actually impact on-the-ground pest control in the future.  By having a better understanding of how termites deal with their dead, the cues they use, and the behaviors those cues elicit, one could imagine how this knowledge could be used to enhance termite exposure to biological or chemical control products in the future.

This research was recently published in the Annals of the Entomological Society of America based on work performed by Jizhe Shi and his colleagues at the University of Kentucky working in Joe Zhou’s lab. Find out more about this research here: Managing Corpses from Different Castes in the Eastern Subterranean Termite

Jim Fredericks, PhD, BCE


 

3 Tips to Take Bug Photos That Entomologists Can Actually Identify

Bug identification is hard, right? That’s why you sometimes need to ask for help from trained entomologists or other qualified experts. Proper insect identification is step one on the path to pest management success for the pest control operator.

Entomologists spend a significant amount of their time identifying insects as a part of their job. Also, as a free consultant to friends, or the random person on facebook they haven’t spoken to in 10 years. If you are a NPMA member and don’t already know, insect identification from a PhD entomologist is a benefit that you have as a member. To help entomologists everywhere get some of the time back they will never see again, follow these tips to be a better bug photographer, even with your phone.

  1. Get close. Repeat after me, GET CLOSE! Most insects are small, some are incredibly teeny-tiny. If the photo is taken several inches away (or feet, I mean really people!?) then you won’t be able to see any detail. Detail is key, so a close photo will help with seeing all the small details that are important.
  2. Keep the insect in focus. Should be a no brainer, right? If the bug is a blurry black dot or not distinguishable then we entomologists will, unfortunately, not be able to perform a miracle identification. Even better, if you can zoom a little and still see somewhat clearly – then you get bonus points.
  3. Take several angles. Just got a picture of the insect back end (abdomen)? An entomologist probably won’t be able to ID without the rest of the insect. Try to get several different angles if you can, especially if your specimen is not alive. Can you take a picture of the top of the insect, the underneath, the face, the legs? It all helps. Sometimes for really specific ID, even a clear picture won’t work unless you can count tiny spines on the legs etc. So, when it gets that crazy it might be time to just mail your bug for someone to look at it under a microscope.

Last but not least, if you get a picture from a customer that you can’t even tell is a bug – trust me, I won’t be able to either.

Brittany Campbell, PhD, BCE