Identification and Treatment of a Storage Mite Infestation in Rats (Rattus norvegicus)

Abstract

Storage mites are environmental pests that commonly invade hay, grain, or stored food. While generally regarded as harmless, they have been reported to elicit allergic reactions in both humans and animals. Although storage mites are considered environmental contaminants, this case report describes an infestation of storage mites on rats (Rattus norvegicus) in a laboratory animal facility. Despite traditional diagnostic methods initially revealing negative results, mites were consistently observed under repeated direct microscopic examinations of the animals. Eventual positive pelage tapes confirmed the presence of the ectoparasites and identified them as mold or cheese mites (Tyrophagus brevicrinatus or Tyrophagus putrescentiae) via Sanger sequencing. To our knowledge, this report is the first to implement permethrin-soaked cotton balls for the successful treatment of mold mites in an entire rat colony. Furthermore, considering the initial negative diagnostic results, this report emphasizes the pelage tape method as highly susceptible to false negatives.

Introduction

Storage mites are environmental pests found in stored food products, including grain, flour, and hay. The Emory University animal care program has a history of intermittently observing storage mites in the rodent diet, and the discoveries generally coincide with warmer, more humid weather in spring and summer, as the mites thrive in these environmental conditions. When found in laboratory animal facilities, the prescribed course of action is to dispose of any infested supplies (such as the feed) and decontaminate the environment (such as sanitizing the housing racks, biosafety cabinets, and food storage bins and changing all animals to newly washed cages). While generally regarded as harmless to animals, storage mites can elicit respiratory and/or dermatologic allergic reactions in both humans and animals,^1–7 causing concern with respect to both occupational health risks and confounds to research studies. A robust review of these pests in the laboratory animal setting, including their biology, potential effects on human and animal health, and management recommendations, has been published.⁸ Of note, it is unlikely that food storage pests are considered in routine pest control measures or colony health monitoring programs, so identification of the tiny mites relies heavily on incidental visualization, such as on food bins or cage lids.

Although storage mites are regarded as environmental pests, this case report describes and documents an infestation of mold mites on animals, specifically rats (Rattus norvegicus), in a laboratory animal facility. The mites were consistently observed on the animals for the duration of the 6-wk investigation. Furthermore, to our knowledge, this report is the first to use permethrin-soaked cotton balls for the successful treatment and eradication of mold mites from the entire rat colony.

Case Summary

In the sole rat housing room within a facility, 2 investigators (henceforth referred to as A.B. and Y.Z.) had census in the room. The 2 groups operated separately in the shared housing space; laboratory members from one group did not interact with the animals belonging to the other group. In July 2024, an investigator (A.B.) notified veterinary staff of ectoparasites visualized on one of their rats during an experimental procedure under a microscope. This A.B. cohort of animals (n = 5) had arrived from an approved vendor only 1 wk prior, and the vendor confirmed a lack of ectoparasites on their most recent health monitoring testing, which included PCR assays and direct examinations. The animal facility is located on the third floor of a hospital building, making a wild rodent incursion unlikely. When asked, animal care staff reported no sightings of wild rodents, and an investigation of the housing room by the pest management company revealed no evidence of wild rodent activity. No construction, renovation, or other major projects were ongoing within the facility, and facility doors were closing and locking appropriately. No special diets were being managed by investigators in this room. An inquiry determined that no laboratory members or animal care staff who frequented this room possessed any pet rodents or snakes. Our routine health monitoring program uses the sentinel-free soiled bedding method, and no fur mites had been detected in this facility via PCR testing for >24 mo.

To confirm the sighting, veterinary staff examined the A.B. cohort of rats but did not visualize any ectoparasites. Six pelage tapes and 2 samples of rodent diet were also collected and examined microscopically, which revealed no ectoparasites. Concurrently, fur swabs were sampled and submitted to a commercial diagnostic laboratory for PCR testing, which came back negative for fur mites, Demodex, and Ornithonyssus. Throughout this time, the investigator continued to report sightings of the ectoparasites on their rats during subsequent experimental procedures. Finally, the investigator was able to capture and submit a clear video of the pests to veterinary staff (Figure 1). Following video confirmation, repeat examinations of anesthetized animals from the A.B. cohort of rats were performed, and ectoparasites were visualized moving along the dorsum of the animals. Repeat pelage tapes were collected at the time of examination that additionally confirmed the presence of mites.

Figure 1.

Still image from the video submitted by the investigator demonstrating the presence of 2 ectoparasites on a rat as seen under a microscope.

Concurrent to the investigation, one rat from the Y.Z. colony developed severe dermatitis highly unusual to the history of this colony. The affected animal was a 10-mo-old, female Brown Norway rat with a 4.0×4.5 cm area of the right scapular region to lateral thorax with moist, sanguinous lesions and dry, thick scabs at deeper lesions. The animal was otherwise bright, alert, and responsive with normal body condition, hydration, posture, and coat. The dermatitis lesions were cleaned with dilute chlorhexidine, hydrolyzed collagen (Collasate spray; PRN Pharmacal, Pensacola, FL) was applied, and nails were trimmed. With the suspicion that an ectoparasite infestation may explain the unusual presence and severity of the dermatitis, an additional pelage tape was sampled directly from this animal. The pelage tape revealed the presence of mites, confirming that both investigators’ colonies within the room were affected.

The video and pelage tape images were sent to multiple experts (that is, parasitologists and entomologists) who all agreed that the ectoparasites had morphologic features consistent with storage mites (for example, grain mite, flour mite, house mite). Therefore, control of the infestation began with environmental decontamination of the entire rat housing room including disposal of the rodent diet, cleaning of the biosafety cabinet, and sanitation of the ventilated racks. Due to the presence of mites in both colonies of animals for several weeks, permethrin-soaked cotton balls were used to treat all of the animals in the room for a duration of 4 wk. Following environmental decontamination and completion of treatment, all subsequent microscopic examinations of the animals performed by veterinary staff revealed no mites, and the outbreak was concluded. Animals in the A.B. colony continued to be monitored by the investigator during experimental procedures with no further reports of ectoparasite sightings.

Pelage tapes positive for mites were submitted to a commercial diagnostic laboratory for Sanger sequencing. While results were not received until after treatment began on the colony based on a presumptive diagnosis, the ectoparasites were ultimately confirmed to be mold or cheese mites (Tyrophagus brevicrinatus or Tyrophagus putrescentiae).

Materials and Methods

Animals and husbandry.

The affected room was the only rat housing room in the entire facility and was located nonadjacent to other rooms housing mice. Only 2 investigators (A.B. and Y.Z.) maintained animals in this room, and census consisted of 3-mo-old, female Wistar rats (n = 5) belonging to A.B. and >9-mo-old, female Brown Norway and Long–Evans rats (n = 63) belonging to Y.Z. All animals were obtained directly from Charles River Laboratories (Frederick, MD).

The Wistar rats were housed in static rat micro-isolator cages (Lab Products, Aberdeen, MD). The Brown Norway and Long–Evans rats were housed in either the aforementioned static cages or ventilated rat Maxi-Miser cages (Thoren Caging Systems, Hazleton, PA). Husbandry methods standard for our institution were performed. All animals were housed on ⅛-in. corncob bedding (Bed-o’Cobs; The Andersons Lab Bedding Products, Maumee, OH), fed irradiated chow (PicoLab rodent diet 20 [5053]; LabDiet, St. Louis, MO), and provided reverse osmosis water in bottles. Bed-r’Nests (The Andersons Lab Bedding Products) were provided as physical enrichment. Per our standard operating procedures, rats are not required to be accessed under a biosafety cabinet; however, a HEPA-filtered class II type A2 biosafety cabinet (NuAire, Plymouth, MN) was available in the room. The room was maintained on a 12-h light/12-h dark cycle with temperatures ranging from 68 to 76 °F (20 to 22 °C) and humidity ranging from 30% to 70% in accordance with the Guide for the Care and Use of Laboratory Animals.⁹ Emory University is accredited by AAALAC International, and all animal work was approved by the Emory University IACUC.

Direct microscopy of animals.

Visual examination for ectoparasites on anesthetized animals was performed using direct microscopy with an Olympus SZX16 binocular zoom stereo microscope (Olympus Corp., Tokyo, Japan). The animals were scanned at 11× magnification, focusing on the dorsum of the animal along the back, neck, and between the ears, the axillary regions, and the inguinal regions.

Direct microscopy of rodent diet.

Visual examination for ectoparasites in samples of rodent diet was performed using direct microscopy with an Accu-Scope 3075 binocular zoom stereo microscope on a coaxial coarse/fine focus LED stand (3075-LED-CF) (Accu-Scope, Commack, NY). Technicians scanned the diet at 4× magnification, which was increased as needed.

Pelage tapes.

Pelage tapes were collected by running a piece of clear adhesive tape partially attached to a glass slide over the dorsum of the animal along the back, neck, and between the ears in the opposite direction to fur growth. Afterward, the tape was fully adhered onto the glass slide and examined for ectoparasites with a Nikon Eclipse E400 biologic microscope (Nikon, Melville, NY) at 20× and 40× magnifications.

Pelt sampling and PCR testing.

Pelt samples were collected using sterile adhesive swabs (VWR, Radnor, PA) and massaging/swabbing the dorsum of the animal along the back, neck, and between the ears in the opposite direction to fur growth. One swab was collected per animal and pooled among 3 groups (A.B. ventilated cages, Y.Z. ventilated cages, and combined static cages). Samples were submitted to Charles River Laboratories for PCR analysis using nucleic acid isolation, PCR processing, and controls as previously described.¹⁰ The testing and interpretation algorithm used by the laboratory is as follows: isolated nucleic acid was screened with proprietary primer and probe sets that target specific rodent pathogens, including fur mites (that is, Myobia, Radfordia, and Myocoptes), Demodex, and Ornithonyssus. To monitor for successful nucleic acid recovery after extraction, and to determine whether there may be evidence of PCR inhibitors, a nucleic acid recovery control assay (sample suitability control) was included with each sample as part of the testing process.

Sanger sequencing.

Extended PCR products were generated by Charles River Laboratories for the pelage tapes using a mite-specific sequencing primer set. PCR products were purified and sequenced using the Sanger method (Genomics Core; Tufts University Medical School, Boston, MA). Sequence data obtained was truncated by removing terminal regions, which included the primers and any undetermined nucleotide bases (Geneious Prime software, version 2024.0.4. https://www.geneious.com). The cleaned consensus sequence was aligned with publicly available sequences in GenBank using the Basic Local Alignment Search Tool (https://blast.ncbi.nlm.nih.gov).

Treatment.

Environmental decontamination began with disposal of all expendable items, including the rodent diet. The empty food bin and extra cages/supplies from the room were sanitized in a BetterBuilt 230 tunnel washer, and racks were sanitized in a BetterBuilt 620 cage and rack washer (BetterBuilt, Delta, BC, Canada). Both washers used Clout as detergent (Pharmacal Research Laboratories, Waterbury, CT). The biosafety cabinet within the room was manually disassembled and cleaned to eliminate debris, particles of bedding, and morsels of rodent diet. Afterward, the cabinet was disinfected using Vimoba (Quip Laboratories, Wilmington, DE) with a contact time of 5 min. All animals were transferred to newly autoclaved cages with autoclaved bedding and water.

Animals were treated with MiteArrest, an insecticidal bedding consisting of cotton (92.6%) and permethrin (7.4%) (EcoHealth, Brookline, MA). By calculation of floor space, 5 cotton balls were added to each Thoren cage, and 10 cotton balls were added to each Lab Products cage. Cotton balls were replaced weekly for a total of 4 wk.

Results

Direct microscopy of animals.

After the investigator captured a video of the pests, veterinary staff anesthetized the remaining 3 rats in the A.B. colony for examination under microscopy. Ectoparasites were visualized moving along the dorsum, at both midshaft and base of the hair, of 2 of the 3 animals.

After the conclusion of treatment, a sampling of rats was examined from both investigator colonies by veterinary staff (n = 4). In addition, animals continued to be regularly examined by the investigator who originally reported the ectoparasites. No ectoparasites have been visualized since the treatment ended.

Direct microscopy of rodent diet.

After the initial report of ectoparasites from the investigator, 2 separate samples of rodent diet were collected from the food bin. Under microscopy, both samples revealed absence of mites.

Pelage tapes.

After the initial report of ectoparasites from the investigator, 6 pelage tapes were collected from the animals (4 samples from the A.B. colony and 2 samples from the Y.Z. colony). All 6 samples revealed absence of mites. Following the video confirmation of ectoparasites, 4 pelage tapes were collected from the A.B. colony, which were all positive for mites. One pelage tape was collected from the Y.Z. animal with severe dermatitis, and it also revealed the presence of mites.

PCR testing.

After the initial report of ectoparasites from the investigator, pelt samples via fur swabs were collected from 6 animals and submitted for PCR testing, which came back negative for fur mites (Myobia, Myocoptes, Radfordia), Demodex, and Ornithonyssus.

Parasitology.

Descriptively, based on the video and pelage tape images, the mites had 8 legs, no palpal claws, and short hairs on the body with long hairs at the distal end of the body. No caudal projecting clasping organs were noted. Sizes approximately ranged from 680 to 840 µm by 470 to 650 µm. The head was ∼200 µm in length. Eggs were oval shaped and approximately 50 to 320 µm by 120 to 140 µm.

Sanger sequencing.

Sanger sequencing was performed on the pelage tapes that were positive for mites. A 473-bp clean consensus sequence dataset was obtained, and Basic Local Alignment Search Tool analysis indicated 100% homology with Tyrophagus spp., supporting morphologic observations. Both Tyrophagus brevicrinatus and Tyrophagus putrescentiae sequences were detected with 100% sequence identity; however, the sequence within the target region cannot differentiate between these 2 Tyrophagus species.

Discussion

This case report describes an infestation of mold mites in a rat housing room. Entry of the mites was presumably via rodent diet, as numerous cages of both investigators were affected, and the initial sighting was reported only 1 wk after the animals arrived from an approved vendor. A novel aspect of this infestation is that the mites were consistently visualized on the animals rather than in the environment throughout the investigation until treatment was concluded.

Treatment of the animals was pursued using permethrin-soaked cotton balls in the entire colony. Permethrin was chosen as the treatment of choice based on its use for environmental decontamination of storage mite infestations^11–13 and Ornithonyssus bacoti infestations in pet and laboratory rodents.^14–20 Four weeks of treatment was administered based on the life cycle of most storage mites and because environmental conditions in the room at that time were generally favorable for this length of life cycle.^1,21–24

Although the life cycle of storage mites can be halted when environmental conditions fall outside their optimal ranges,^8,21 it was not possible to adjust the temperature or humidity of the room sufficiently outside the parameters while staying within the Guide for the Care and Use of Laboratory Animals requirements for the animals. Therefore, the animals were treated directly, at the cage level. A report of the temperature and relative humidity within the affected room indicated no significant changes during the period of this infestation (72.20 ± 0.25 °F [22.3 ± 0.45 °C]; 63.05 ± 0.68%), affirming the conclusion that the mites were eradicated from the permethrin treatment rather than from environmental alterations or fluctuations.

Tyrophagus spp. are common pests of stored products with an affinity for protein- and fat-rich content, granting them colloquial names such as the cheese mite, ham mite, or copra mite.¹ Notably, this genus of mites causes the well-described copra itch experienced in humans.²⁵ Ideal conditions for Tyrophagus spp. are temperatures >86 °F (30 °C) and humidity >85%, under which their life cycle can be completed in <3 wk.²⁶

In some mite species of the order Acarina, a special deutonymph stage, known as the hypopus,^1,27,28 may develop when environmental conditions are unfavorable, such as when there is a lack of food, overcrowding, or desiccation. In this stage, suckers develop on the underside of the mites, aiding in their dispersal by allowing them to attach to animals or other insects. The hypopus may exist for several months with no nutrition.²⁹ It does not grow and is highly stable, making it resistant to typical insecticides. Interestingly, the Tyrophagus genus has lost its ability to form hypopi,²⁷ but due to the implications for control measures, this unique form should be considered when encountering other common storage mites of laboratory animal facilities, such as the grain mite (Acarus siro). This stage could also contribute to the resurgence of mite infestations after chemical treatments seem effective.

Another aspect of note in this case report is the high percentage of false negatives obtained from the examination of grain samples and pelage tapes. At the initial sampling timepoint, 2/2 (100%) grain samples and 6/6 (100%) pelage tapes revealed absence of ectoparasites even though mites were observed directly on the animals. Therefore, these techniques should be regarded as highly prone to false negative results.

This report describes an unusual presentation of an environmental pest, Tyrophagus spp., on rats. After multiple rounds of diagnostic testing, the mite was definitively identified, and successful eradication was performed using permethrin-soaked cotton balls. Facilities with a history of storage mites should consider the ectoparasites as a differential diagnosis for allergic dermatitis in animals, and there may be a need for repeated diagnostics and treatment at the cage level if environmental parameters at the room level cannot be sufficiently adjusted to control the infestation.

Conflict of Interest

The authors have no conflicts of interest to declare.

Funding

This work was internally funded.