Rearing Phoretic Mites Associated with Wood-Boring Insects

Summary

Here, we present a protocol to establish and maintain laboratory cultures of phoretic mites associated with wood-boring insects, including both predatory and fungivorous species, using a rearing unit with customizable food sources and scalable methods for experimental or applied research.

Abstract

Phoretic mites are common associates of a wide range of wood-boring insects, such as Coleoptera: Curculionidae and Cerambycidae. Phoretic mites are among the few groups of organisms capable of accessing their hosts' breeding sites. These organisms are minute, establishing close associations and affinity for biotic and abiotic conditions within wood-boring insects' galleries. Their presence within the galleries may exert a significant influence on beetle behavior and reproduction. Some species may act as natural antagonists of the beetles and their mutualistic fungi, disrupting symbiotic relationships that guarantee their success. These characteristics make phoretic mites promising candidates for biological control programs targeting wood-boring pests. However, reproducible methods for rearing these mites are needed to study their basic biology and ecology and assess their potential as biocontrol agents. Despite their importance, there are currently no detailed protocols for maintaining phoretic mite populations in captivity. This study presents a practical and accessible approach for rearing two groups of phoretic mites, fungivores/detritivores (Acari: Astigmata) (i.e., Histiogaster arborsignis) and predators (Acari: Mesostigmata) (i.e., Proctolaelaps spp.). We propose using rolled barley grains and nematodes as nutritional sources for these mites, respectively. These food sources are practical and widely accessible, requiring minimal maintenance while offering a standardized diet tailored to the nutritional requirements of the target mite groups. A plaster-charcoal-based substrate is used as a humidity-regulating matrix. This combination provides both food resources and a suitable microhabitat for long-term maintenance and reproduction of these mites. The method described is scalable, enabling researchers to expand rearing units in proportion to colony size and experimental requirements. This protocol offers a critical tool for advancing research into the ecological roles of phoretic mites. This experimentally validated system ensures long-term colony viability for several months and multiple generations, while remaining adaptable for rearing other mite taxa under laboratory conditions.

Introduction

Many wood-boring insects complete most of their life cycle inside trees, where they feed and often act as vectors of plant pathogens, posing serious risks to agriculture and forest ecosystems^1,2. Among the most damaging groups are invasive beetles in the families Buprestidae, Cerambycidae, Bostrichidae, and Curculionidae, which have been linked to extensive tree mortality and economic losses worldwide³. Florida hosts over 35 ambrosia beetle species (Coleoptera: Curculionidae), with at least 15 known to infest avocado trees^4,5. Common species in orchards include Xyleborinus saxesenii Ratzeburg, Xyleborus affinis (Eichhoff), X. bispinatus Eichhoff, X. ferrugineus (Fabricius), X. volvulus (Fabricius), X. gracilis (Eichhoff), and Xylosandrus crassiusculus (Motschulsky), often co-occurring in the same tree^4,6. These beetles are damaging primarily because they vector symbiotic fungi, some of which are pathogenic to plants⁷. The most serious pathogen is Harringtonia lauricola, the causal agent of laurel wilt, introduced to the U.S. with its primary vector, Xyleborus glabratus Eichhoff, around 2002^8,9. Laurel wilt has devastated Florida's avocado industry and native Lauraceae, killing over half a billion trees^10,11. Moreover, H. lauricola now occurs in association with multiple ambrosia beetle species, which function as alternative vectors in avocado systems where X. glabratus is rare^4,12.

Mites (Acari) are highly diverse arthropods occupying a wide range of ecological niches. Among them, several groups are increasingly recognized for their potential as biological control agents due to their predatory behavior, high reproductive rates, and ability to thrive in various environments, including soil, plant surfaces, and stored products. Their ability to suppress pest outbreaks in greenhouses, nurseries, and agricultural fields underscores their importance in integrated pest management (IPM) programs^13,14. For example, the family Phytoseiidae includes several species of considerable importance in biological control, notably Neoseiulus californicus Berlese and Phytoseiulus persimilis Athias-Henriot, which are extensively utilized for the suppression of spider mites (Acari: Tetranychidae) and thrips in greenhouse and field crops¹⁵. Likewise, the laelapid mite Stratiolaelaps scimitus is a biological control agent targeting soil-dwelling pests such as fungus gnat larvae and thrips pupae and is broadly incorporated into integrated pest management programs¹⁶. Phoretic mites represent a promising group in this context. These mites rely on temporary associations with their hosts for dispersal and are frequently found within the galleries or brood chambers of wood-boring insects such as bark and ambrosia beetles^17,18,19.

The diversity of phoretic mites extends to both morphology and ecology. Some exhibit fungivorous and detritivorous feeding behavior, while others are predatory. Considering their closeness to their hosts, phoretic mites have the potential to suppress insect populations directly by predating on eggs and larvae, by physically impairing them, or indirectly, by disrupting symbiotic associations of wood-boring insects^20,21,22,23.

Despite the growing interest in mites and their potential biological control agents, the development of rearing protocols for them remains limited. Some studies have reported methods for culturing free-living predatory mites (Acari: Mesostigmata)^24,25; however, the descriptions often lack instructions for the initial establishment of mite colonies, posing a challenge for new acarologists. Currently, no established protocols are available for maintaining long-term laboratory colonies or rearing phoretic mites for experimental use. The protocol presented in this study addresses this gap by offering a practical, scalable, and reproducible method for rearing both predatory (i.e., Mesostigmata) and fungivore/detritivore (i.e., Astigmata) mites. Although the protocol is described in the context of phoretic mites, it is broadly applicable to other mites and can be readily adapted to suit diverse research objectives. For instance, Astigmata mites reared using this method can serve as factitious prey for predatory mites^26,27.

This study aimed to develop and evaluate a standardized rearing protocol for phoretic mites, under the hypothesis that a broadly applicable substrate and diet combination can support sustained survival and reproduction across diverse mite taxa (Mesostigmata and Astigmata).

Protocol

The protocol described here is an upgraded adaptation of previous descriptions of mass-rearing techniques for soil arthropods^24,25. It also includes a new method for mass-rearing fungivore/detritivore phoretic mites (i.e., Astigmata) using autoclavable plastic bags. The reagents and the equipment used are listed in the Table of Materials.

1. Preparation of the rearing unit substrate

1. When starting a culture with 1-5 mites, use a 5 mL screw-on lid jar to facilitate mite visualization during colony establishment. To know the starting jar size, calculate approximately 20 mites per 30 mL container volume.
   NOTE: For example, when starting a culture with 5-20 mites, use a 30 mL or smaller screw-on lid clear jar. When starting a culture with 20-40 mites, utilize the next jar size (60 mL).
2. Prepare a dry mix of 90 g of plaster of Paris and 10 g of powdered activated charcoal (9:1) in a 500 mL autoclavable beaker. Stir the powder until fully mixed before autoclaving.
3. Cover the beaker with aluminum foil to prevent water from entering the container during the autoclaving process.
4. Prepare a mixture of organic bark pieces and Sphagnum moss (2:1) in an autoclavable beaker. Prepare enough to cover about half of the rearing container.
   NOTE: For example, when preparing a 120 mL container, prepare about 60 mL of the substrate.
5. Cover the beaker with aluminum foil to prevent water from entering the container during the autoclaving process.
6. Pour 500 mL of distilled water into an autoclavable beaker.
7. Cover the beaker with aluminum foil.
8. Autoclave the dry mix, the bark-moss mixture, and the distilled water in separate covered beakers. Set the autoclave for 30 min at 121 °C and 15 psi, with a 30-min drying time.
9. Allow the materials to cool down to room temperature (25 °C) for about 1 h. Leave all materials covered with foil while cooling.
10. Once cool, pour 50 mL of autoclaved distilled water into the activated charcoal-plaster mix at a time, mixing the blend every time more water is added. Repeat the process until a creamy consistency is achieved (the ideal consistency is similar to mayonnaise).
11. Immediately pour the mixture into the bottom of the jar and tap it gently a few times against a flat surface to eliminate the air bubbles. Pour enough to create a 1 cm layer of the mixture.
    NOTE: If using a jar smaller than 30 mL, adjust the thickness of the layer to about 25% of the jar height.
12. Wait for the mixture to dry completely (about 6 h) (Figure 1A,B).
13. Add distilled water to the bottom layer until it turns a darker gray. Add enough water to saturate this layer without making the surface wet. If an excessive amount of water is poured, use a piece of paper towel to absorb the excess water.
14. Add the autoclaved mixture of organic bark pieces and Sphagnum moss (2:1) to the rearing unit. Add enough to cover half or less of the rearing unit (Figure 1C).
    NOTE: If starting a culture with 1-5 mites, add a single small piece of bark and/or a strand of Sphagnum moss to facilitate mite visualization during colony establishment (Figure 2A).

2. Preparation of a ventilated lid

1. Drill a hole in the center of the jar lid. For a 30 mL jar, drill a 1 cm diameter hole. For larger containers, increase the size of the hole at a rate of about 2 mm diameter for every 30 mL increase in the container's volume (i.e., 1.2 mm diameter hole for a 60 mL container, 1.4 mm for a 90 mL container, etc.).
   NOTE: When using a jar smaller than 30 mL, it is not recommended to drill a hole in the lid, as the substrate will dry out quickly (Figure 2B). Ensure that the container is opened every 2 days for air exchange.
2. Use hot glue to affix a mite-proof mesh (80 µm aperture or less) on top of the hole. Apply the hot glue around the hole on both sides of the lid to ensure that all crevices are covered (Figure 1D).

3. Food source and phoretic mite transfer

1. Offer small patches (about 20% of the surface) of prey/food items separately in the rearing unit.
   NOTE: Determine the food source based on the feeding habits of the mite being reared. For predatory mites (i.e., Mesostigmata), offer nematodes and/or larvae and eggs of Astigmata mites (i.e., Aleuroglyphus ovatus orThyreophagus entomophagus)²⁷. The most accessible and easy-to-culture nematode to feed predatory mites is microworms (Panagrellus spp.).(Nematoda: Panagrolaimidae), readily available commercially for use as a food source for fish fry. Alternatively, utilize commercially available entomopathogenic nematodes, such as Steinernema spp. and Heterorhabditis spp.²⁸. For fungivores/detritivores (i.e., Astigmata), offer rolled barley grains and rolled oats.
2. Transfer the mites to the rearing unit using a fine paintbrush.
3. Label the cultures. Write scientific and common names (when available), date, location of collection, and host.
4. After 24 h, observe the mites under the stereomicroscope to determine what prey/food item is preferred by them.
5. Keep the cultures at a temperature range of 25-27 °C in complete darkness.

4. Host and phoretic mite collection

1. Choose and locate the phoretic hosts. Many wood-boring beetles and other insects serve as phoretic hosts of mites. Scout areas and specific ecological niches occupied by the phoretic host.
2. Collect the hosts employing methods that allow the phoretic mites to remain alive after collection, such as baiting, trapping, net collection, and/or handpicking. Avoid using killing agents.
   NOTE: The collection method will depend on the phoretic host. For example, bark and ambrosia beetles can be collected using Lindgren funnel traps, by manually excavating logs, or by placing infested logs in containers that serve as emergence chambers²⁹.
3. Transfer the phoretic hosts to the laboratory.
4. Use a stereomicroscope (40-200x) to locate the phoretic mites on the host's body.
   NOTE: Phoretic mites are commonly attached to beetle wings (elytra), soft cuticle (i.e., between leg and body segments), thorax, and under the elytra.
5. After preparing the rearing unit (following step 2, step 3 and step 4), extract the mites carefully from the hosts' body using a fine paintbrush.
6. Examine the collected individuals under a stereomicroscope (40-200×) to verify that all mites added to the rearing unit are morphologically uniform. If multiple morphotypes are detected, separate them and establish individual rearing units for each distinct morphotype.
7. Transfer the phoretic mites to the rearing unit using a fine paint brush.

5. Culture maintenance

1. Observe the cultures under a stereomicroscope (40-200×) every 2 days. Check for signs of contamination with sporophytes (mold), other mites, and bacteria.
2. Manually eliminate contamination using a sterilized tool like forceps. Small mold growths can be managed by gently applying 70% EtOH directly to the hyphae with a paint brush. Eliminate any unrecognized mites.
   NOTE: Some mite life stages look morphologically distinct from others. For example, many astigmatid mites will produce phoretic deutonymphs (hypopi) that do not bear resemblance to other life stages. Research the literature for morphological differences between the life stages of the mite being reared.
3. Replenish the food source. Add just enough food to be consumed before the next culture maintenance to avoid fouling the rearing unit.
4. Add distilled water to the bottom of the experimental unit to keep the humidity high (>75%). Use a pipette to add water in 1 mL increments. Add enough water to saturate this layer without making the surface wet (shiny). If the surface becomes excessively moist, use a paper towel to absorb the excess water until the surface becomes opaque.
   NOTE: As the layer absorbs water, it will turn a darker gray.
5. Once a week, gently shake the rearing unit to avoid substrate compaction. This step is important for cultures being kept in containers above 500 mL.

6. Scaling up predatory phoretic mite cultures

1. After 2-4 weeks of culture establishment, or when the culture is saturated with mites, prepare a larger-sized jar equipped with prey following the steps described above. To know the size of the jar needed, estimate the number of mites being transferred and utilize the rule of 20 mites per 30 mL.
   NOTE: When the culture is saturated with mites, they will start crawling on top of the substrate, as well as on the lid. This is generally the time to upscale the culture to a larger container.
2. Transfer all the substrate with mites from the old jar to the new one.
3. Turn the old jar upside down above the new jar and tap the bottom to dislodge mites from the old arena onto the new rearing unit.
4. If a substantial number of mites are left in the old jar, manually transfer them using a fine paint brush.

7. Scaling up fungivores/detritivores cultures to mass rearing plastic bags

NOTE: As fungivore/detritivore mites consume the grains, their frass and decaying organic matter can quickly foul the rearing substrate, making the conditions harmful for the mites. Therefore, small rearing units, up to a 120 mL jar, are recommended to maintain small colonies of fungivorous mites. For scaling up or mass rearing fungivorous mites, autoclavable bags with rolled barley are recommended.

1. Add 100 g of rolled barley or oats to a 19 cm x 19 cm x 32 cm filtered autoclavable bag (commonly used for mushroom cultivation) (Figure 3).
2. Add 25 mL of distilled water to the grains.
3. Close the bag loosely so that it does not burst in the autoclave.
4. Autoclave the bag using the Liquid 30 cycle (121 °C).
5. Wait for the grains to cool down completely before introducing mites.
   NOTE: The grains should be soft and moist after being autoclaved. The initial amount of water might depend on grain type and brand, due to differences in the way they are processed. The volume of water added to the grains may be adjusted by increasing or decreasing it by 5-10 mL from the protocol recommendation (25 mL). For example, if the grains look completely dry, add an extra 5-10 mL of water before autoclaving. If visible water is present at the bottom of the bag, reduce the amount of water added before autoclaving by 5-10 mL.
6. Transfer the mites from the rearing jar to the bag. Use a small teaspoon or equivalent to scoop a mix of all life stages from the surface. Transfer only the mites with the grain substrate. Do not transfer the pieces of bark and Sphagnum moss, as they are sources of mold contamination.
7. Seal the bag using a heat plastic bag sealer.
8. Keep the cultures at a temperature range of 25-27 °C, preferably in the dark.

8. Fungivores/detritivores culture maintenance in plastic bags

1. As the mites consume the grains, they are turned into powder and tend to compact. Agitate the bags twice a week to ensure that the food substrate is aerated and mixed in.
2. When most (>75%) of the grains have been consumed or if there is contamination with fungi or bacteria, prepare a new bag following step 8.
3. Transfer the mites to the new bag. Ensure transferring the powdered substrate, as it will have the highest concentration of mites.
   NOTE: The time mites should be left in the bag before being transferred to a new one will depend on the mite species in question, the initial number of mites introduced, and the temperature they are kept at. In most cases, the bags will last for about three weeks before they must be replaced.

9. Monthly colony inspection

1. Once a month, prepare microscope slides of 5-10 adult mites in the colony using Hoyer's medium³⁰.
2. Check the specimens under the microscope (400-1000x) to ensure that the mite species being reared has not been replaced by a contaminant.

Results

The protocol described in this manuscript effectively supported the growth and reproduction of two predatory mite species, Lasioseius safroi (Mesostigmata: Blattisociidae) and Proctolaelaps sp. (Mesostigmata: Melicharidae), and a fungivore/detritivore species, Histiogaster arborsignis (Astigmata: Acaridae), under laboratory conditions. Ten colonies (replicates) were prepared for each mite species. Each colony was initiated with five individuals kept in 5 mL arenas (as described in steps 2, 3, 4...

Discussion

The present study demonstrates that the proposed rearing protocol effectively supported the survival and reproduction of both predatory mites (Lasioseius safroi and Proctolaelaps sp.) and a fungivorous/detritivorous species (Histiogaster arborsignis) under controlled laboratory conditions. Substantial population increases were recorded over the 14-day period, with mean colony sizes exceeding 200 individuals following the transition from mL to 120 mL arenas. These results confirm the robustness ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
5 mL Clear Containers With Lid	Amazon - Medsuo	‎Y-ZS-11-299
Detail Thin Paint Brush Set 6 pcs	Amazon - Transon	SJGGX6
Everyday Aluminum Foil	Reynolds Wrap	https://www.reynoldsbrands.com/products/aluminum-foil/standard-foil?gad_source=1&gad_campaignid=19 457576830&gbraid=0AAAAADmFV MecGX9nzTc8tAkfL1_tRwqyv& gclid=Cj0KCQjwhafEBhCcARIs AEGZEKLB5fdLiDNOqPXlxtZA -ZlGEYQjzLX5w7KmrTzF6-_ q8sabcwroobIaAlmPEALw_wcB
Impulse Heat Bag Sealer	Amazon - runruii	619438296390
Multi Filter Strips Autoclavable Bag	Amazon - Microsac bags	B076C2KTMD
Organic Coconut Activated Charcoal (Powder) Food Grade	Amazon - Belle Chemical	AC - 01
Plaster of Paris (Powder)	DAP	10318
Repti Bark	Zoo Med	RB4
Rolled Barley Flakes	Amazon - Mulberry Lane Farms	B015EURFTY
Rolled Oats	Quaker	30000562314
Sphagnum Moss	Amazon - Riare	B0B1TCJQ6R
Stainless Steel Woven Wire 200 Mesh	TIMESETL	TXJ-484-US
Steris Amsco Lab 250	Steris	RSLAB250
Walter Worm - Panagrellus silusioides Culture	Amazon - Josh's Frogs	B08XY3LM46
Wide Mouth Plastic Jars Round Clear with Black Lids	Amazon - Eternal Moment	734172973182	30 to 250 mL available