David Evans Walter
Heather C. Proctor
Soil and litter habitats have become recognized as important repositories for biodiversity. They are also dominated by some of the smallest and most difficult to identify of all animals · the mites (Arachnida: Acari). This publication represents the authors· attempts to make identification of soil mites less daunting, and to provide up-to-date diagnoses and classifications at the subordinal and family levels.
This volume contains 5 interactive keys for the identification of 101 families of soil mites and more than 60 higher-level taxa of soil microarthropods and related groups (Onychophora, Tardigrada). Although they were constructed with the soil fauna of Australia in mind, the keys cover most classes and orders of terrestrial arthropods and most families of soil-litter inhabiting mites (except Oribatida and Astigmata) at the world-level. Those interested in identification of oribatid mites (Oribatida = Cryptostigmata) below the cohort level should consult Hunt et al. (1998).
These interactive keys are designed to help beginning students to identify the major taxa of soil-dwelling arthropods and to aid more advanced researchers to identify families of soil mites. Each taxon page has information on morphology, similar species, ecology, and critical references. To accommodate those with little knowledge about mites we have used relatively non-technical characters when possible. Each character state is illustrated and explained in an accompanying html file. Additional information on the morphology, behaviour and ecology of the mites is presented in the General Information sections of each key.
The keys were created in Lucid Professional Version 2.0 (http://www.lucidcentral.com/), software that allows the construction of image-rich, html-linked diagnostic tools. Major taxonomic groups of soil-inhabiting arthropods are presented in the first key: Classes and Orders of Microarthropods in Soil (31 classes and orders). The major taxonomic groupings of mites · including Oribatida - are distinguished in the second key: Mites in Soil · Orders, Suborders, & Cohorts (31 suborders and cohorts). Members of the soil Prostigmata (Trombidiformes) can be identified using the key Families of Prostigmata in Soil (43 families). Mesostigmata, Holothyrida, and Ixodida are identified using Families of Parasitiformes in Soil (46 families). Members of the Endeostigmata and Sphaerolichida can be keyed using Endeostigmata and Sphaerolichida (12 families).
The remainder of this booklet deals with the following topics: classification of the Acari; differences between types of identification tools; how to use the enclosed interactive keys; how to extract, preserve, and prepare mites (and other microarthropods) for identification and how to study living mites.
Higher-level Classification of Mites
Mites have been traditionally treated as a Subclass (Acari or Acarina) of the Class Arachnida, and our keys follow this tradition; however, in some recent studies the taxonomic rank of mites has been raised. The closest relatives of mites appear to be a small group of arachnids called the Ricinuleida (no common name). The Acari + Ricinuleida comprise the Acarinomorpha. Shared derived characters supporting this taxon include a hexapod larval stage and a distinct anterior body unit called the capitulum or gnathosoma. Within the Acarinomorpha are four orders, the Ricinuleida and three orders of mites: Opilioacariformes (= Tetrastigmata, Notostigmata, Opilioacarida), Parasitiformes (ticks, holothyrans, and mesostigmatans), and Acariformes (endeostigmatans, oribatids, astigmatans, sphaerolichids, and prostigmatans). The Opilioacariformes and Parasitiformes appear to be closely related, but convincing synapomorphies have yet to be demonstrated. Some authors group these two orders into a superorder (Anactinotrichida) based on the plesiomorphic lack of actinochitin (a substance that is refractive under polarized light) in their setae and oppose it to the superorder Actinotrichida (=Acariformes) whose members have actinochitin-filled setae.
The three major lineages of mites have been recognized since the seminal studies of Grandjean in the 1930's, but well defined subgroups have emerged more slowly. The Parasitiformes has three distinct suborders: Ixodida (ticks), Holothyrida, and Mesostigmata (= Gamasina). The Mesostigmata is further divided into what are traditionally called ·cohorts· (taxa holding the rank of infrasuborders). Fourteen of these cohorts occur in soil and are differentiated in Mites in Soil · Orders, Suborders, & Cohorts. Forty-six families of parasitiformans that live in soil are keyed in Families of Parasitiformes in Soil.
The order Acariformes contains about 80% of the 50,000+ described species of mites, and is split into a morass of subdivisions. The currently recognized suborders include Oribatida (= Cryptostigmata, Oribatei), Astigmata (= Acaridida), Endeostigmata, Sphaerolichida, and Prostigmata (= Actinedida). To further complicate things, the Astigmata and most of the Endeostigmata should be grouped with the Oribatida. The group formed by this union is called the Sarcoptiformes. Likewise, the remaining two families of the Endeostigmata (comprising the Sphaerolichida) should be grouped with the Prostigmata, thereby producing the Trombidiformes. In Mites in Soil · Orders, Suborders, & Cohorts, the Sphaerolichida, 6 cohorts of Prostigmata, the Astigmata, the unusual dispersal morphs of the Astigmata (Astigmatan hypopi), the Endeostigmata, and 7 cohorts of Oribatida are differentiated. In Endeostigmata and Sphaerolichida, 12 families are identified. In Families of Prostigmata in Soil, 43 families are identified.
Dichotomous Keys & Other Identification Tools
The most common identification tool is the dichotomous key:
· ·Dichotomous· means split into two, and refers to the paired phrases opposed in each unit of the key. More than two phrases opposed is considered bad form.
· ·Key· refers to a solution to what is unknown or mysterious and has been used to refer to these identification tools since about the mid-1800·s (Oxford English Dictionary)
· Dichotomous keys are composed of couplets (diminutive of ·couple·), i.e. a pair of statements that provide contrasting characters. Each statement is called a ·lug·, perhaps for the same reason as are the pair of flaps that cover one·s ears on some hats.
· A series of couplets are arranged so that making choices among the lugs moves one along a path through the key towards an identification.
Dichotomous Key to Distinguish Mites from Other Soil Arthropods
1. Body composed of 2 or 3 regions, including a distinct head that usually bears 1-2 pairs of antennae and often has eyes (sometimes compound), opposed mandibles (often internal or modified into stylet-like structures), and 1-2 pairs of small, segmented palps; head followed by a trunk with numerous (>3) segments each bearing 1-2 pairs of legs or by a distinct thorax with 3 pairs of legs and sometimes with 1-2 pairs of wings (anterior pairs sometimes shell-like) and a segmented abdomen with or without limb-like structures; 2-3 terminal cerci (segmented ·tails·) often present........................................................................................ Mandibulata
- Body composed of 1 or 2 regions, neither of which is a head (although a head-like capitulum may be present); antennae never present (although antenna-like pedipalps sometimes present); compound eyes and opposed mandibles never present; 1-4 pairs of simple ocelli may be present; anterior body region combining feeding and locomotory functions and usually bearing 5-6 pairs of limbs: pincer-like, fang-like or stylet-like chelicerae, pedipalps, and 3-4 pairs of walking legs; posterior region without leg-like limbs, although segmented spinnerets sometimes present, and sometimes terminating in a single flagellum or sting ......................... Arachnida (2)
2. Abdomen without distinct segmentation.................................................................. 3
- Abdomen, at least ventrally, covered in segmentally arranged plates...................................................................................... most Arachnida except spiders and mites
3. With narrow pedicle (·waist·) joining front and rear body regions; chelicerae fang-like with large rectangular bases; abdomen with spinnerets.......................... Araneae
- Without pedicle between front and rear body regions; chelicerae various; abdomen without spinnerets............................................................................................. Acari
· Notice that the lugs in each couplet are constructed more or less in parallel, i.e. each choice given in the first lug has an alternative in the second lug. Shortcuts, such as ·not as above·, are considered poor form (and confusing for the user).
· Usually, the more characters given in a couplet, the more likely the user is to be able to identify an organism. For example, if the abdomen of the specimen being keyed is missing or segmentation is indistinct, then couplet 2 above is of no help.
· If you aren·t sure which lug to follow, then mark where you have a problem and try both of the resulting paths.
· Although the structure of the key above is one that is commonly used, other forms exist. For example, in much of the European literature, couplets may be split into different sections of the key with each number followed by a number in parentheses. If characters don·t fit the specimen, then you move to the number in parentheses. If the characters do fit, you go to the next line until an identification has been made. For example:
European-Style Dichotomous Key to Suborders of Soil Mites based on Adult Females
1 (2). Subcapitulum with beak covered with retrorse denticles........................ Ixodida
2 (1). Subcapitulum without retrorse denticles.
3 (4). With 4 pairs of dorsolateral stigmata, without peritremes; tritosternum present and composed of two finger-like processes; rutella present and strongly denticulate........................................................................................................... Opilioacarida
4 (3). With a single pair of tracheal openings or stigmata absent; tritosternum present or absent; rutella present or absent.
5 (10). Prodorsum with 1-2 pairs of trichobothria.
6 (9). Rutella present and well developed; prodorsum usually with 5-6 pairs of setae, including 1-2 pairs of trichobothria.
7 (8) Body covered in well sclerotized plates; notogaster with a maximum of 16 pairs of setae; prodorsum with 1 pair of trichobothria............................................ Oribatida
8 (7). Soft-bodied; notogaster indistinct, usually with >16 pairs of setae, sometimes hypertrichous.................................................................................... Endeostigmata
9 (6). Rutella absent; prodorsum usually with 4 (rarely 5) or fewer pairs of setae or hypertrichous.............................................................................. Prostigmata (pars)
10 (5). Prodorsum without trichobothria.
11 (14). Subcapitulum with a pair of horn-like corniculi; chelicerae 3-segmented; stigmatal opening lateral, above freely articulated coxae
12 (13). Subcapitulum with 5 or more pairs of setae; each chelicera with 2 setae; anal valves with 2 or more pairs of setae....................................................... Holothyrida
13 (12). Subcapitulum with 4 pairs of setae; each chelicera with a single seta; anal valves with 0-1 pairs of setae.............................................................. Mesostigmata
14 (11). Corniculi absent; chelicerae 2-segmented; stigmatal openings between cheliceral bases or on prodorsum; coxae of legs fused to body wall.
15 (16). Palps with 2 free segments; leg tarsi with a single claw; large dorsolateral opisthosomal gland usually present.......................................................... Astigmata
16 (15). Palps with 3-5 free segments; leg tarsi usually with 2-3 claws; opisthosomal gland absent............................................................................... Prostigmata (pars)
Both styles of dichotomous keys have serious drawbacks: one must start at the first couplet regardless of whether it presents the best or most readily seen characters; if the structures described in a couplet are missing, identification is stymied; and illustrations and definitions of structures are usually separated in space from where they are mentioned in the key. Interactive computer-based keys overcome these problems.
How to Use the Interactive Keys to Soil Mites and Other Microarthropods
This is a quick overview of how to use LucID keys in the LucID Player program. More detailed information on keying strategies can be accessed under Help/Contents in the top menu of the Player. Likewise, there is taxon-specific information for each key under Help/About this Key and under General Information (click the blue ball with the ·i·). We strongly recommend that you read this information prior to your first use of a key.
To use these keys to identify mites, you usually will need slide-mounted specimens (see below) and a compound microscope. You should be able to identify most non-mite microarthropods using a dissecting microscope and Classes and Orders of Microarthropods in Soil. If you wish to key a mite to family and cannot immediately recognize to which order it belongs, you should use either the dichotomous keys in this booklet or the interactive key Mites in Soil · Orders, Suborders, & Cohorts. Oribatid mites (including Astigmata) are not keyed beyond cohort level (see Hunt et al. 1988 for species-level identification of the Australian fauna), but most members of the Parasitiformes, Prostigmata, Endeostigmata, and Sphaerolichida can be keyed to family by using the appropriate key on this CD.
When using a LucID interactive key, you will see four windows on the computer screen. The upper left is a list of all characters. Each character has two or more character states. The upper right window is the repository for selected states. The lower left is the repository of discarded taxa, and the lower right for taxa that are still eligible. You may start your identification with any character in the key. Until you become confident in your anatomical expertise, you should retain the default option ·Allow Misinterpretations· in the Key menu. Certain common mistakes are accepted as ·correct· answers when Allow Misinterpretations is operating. You can uncheck this function only when you first start (or restart) the key.
There are two windows for each character state: an image and a note window (the latter often with additional images available through hyperlinks). Click on the coloured button to the left of the character to view the character state image window, and on the line to the right of the button to avoid the images. Notes about character states are present in both modes, but if using the image window, you must drag the thumbnail over the projector icon to view the note. Control-click enlarges an image (or drag it over the projector icon). Double-click on the state you wish to select (you may choose more than one state from a character) and the next character state comes up. Selected states will move to the upper right quadrant, and any taxa lacking those states will move to the lower left quadrant, but you will have to shut the character state window to see these changes. Pressing the ·Best· button at the top of the screen rearranges the characters so that the one most useful in separating the remaining taxa is moved to the top of the list. Note, however, that ·best· doesn·t mean ·easiest to see·!
Information about taxa can be viewed at any time by clicking on the coloured button next to the taxon name. You can choose to read about the taxon or to look at images. Images can also be accessed within the text ·Overview· section, which has this information:
· classification and diagnosis
· list of similar taxa
· ecology and distribution
· list of taxa known from Australia
· references
Hyperlinks to images are present in some of these sections. If you come to an impasse and cannot assign your mite to a single taxon, then look at diagnoses of the remaining candidate taxa. We have included more detailed morphological information in this section. You can also use the Similarity/Difference function or the ·Bingo· button (which lists character states that · if present · would give you an instant identification of the remaining taxa).
Extracting, Preparing and Mounting Mites
Although the following sections are tailored to the Acari, most methods will work well for other microarthropods.
Extracting Mites from Substrates
(a) Funnel extraction. The easiest way to collect large numbers of mites and other microarthropods from organic substrates is to use a funnel or canister extractor. In this simple process, a sample is suspended over a funnel, a humidity gradient is established, and the escaping animals are funnelled into a preservative or live-collecting vial. Heat is used to establish a humidity gradient, and Antonio Berlese developed the first extraction funnel using a hot water bath as a heat source. A later modification involved replacing the water bath with a light bulb. Various versions of the Berlese funnel (or more pedantically, Tullgren funnel, named after the first worker to use light bulbs) are most often used for extracting mites from substrates, including soil, litter, nests, and vegetation. Funnel extractors can be as simple as a goose-neck lamp, a piece of screen, and a plastic funnel supported by an inverted cardboard box, or they can be as intricate as your budget allows. We recommend that anyone undertaking a quantitative sampling program review the recent literature.
When electricity is not available, heavier than air volatiles (e.g. naphthalene crystals sprinkled on the surface of the sample) have been used. Such chemical extraction techniques require good ventilation and an understanding of the toxicity of the chemical. Drying samples in the sun will also extract mites, although this tends to be a slow procedure and is impractical in wet climates. Norton and Kethley (1988) have designed a portable funnel system for use in sun-drying. Painting the reflector cover black may increase its efficiency.
The walls of a Berlese funnel should be steep-sided and the screen mesh should be large enough to let through the largest mite. A layer of muslin under the sample will reduce the amount of dirt falling through. Open funnels may attract flying insects (and their mites), which will add spurious specimens to a collection, but closed funnels need enough ventilation to allow the sample to dry. Funnels work by using heat to dry a sample from top to bottom. Mites must be able to follow the humidity gradient that develops through the samples to the screen and thence tumble into the funnel and collecting vial. DO NOT put too much material in a funnel. If you put in too much, then the centre of the sample may remain moist and mites will follow the humidity gradient and die in the sample. Also, thick, wet samples can become enmeshed in a fungal mat that prevents animals from escaping.
Heat and photonegative reactions are minor factors in funnel extraction and it is important to remember that the humidity gradient is primary. For example, funnels can be used to extract mites from bark and twigs, but many will follow the humidity gradient deep within the sample, where they will die, rather than tumbling into the collecting vial. Mesic habits, e.g. moist (but not dripping wet) soil, leaf litter and compost, are most appropriate for a funnel extraction, but drier organic materials like nests can also be productive if they are composed of relatively fine litter.
Mites living in dry grassland or desert soils, deep mineral soils, or coarse woody material (e.g. bark) are much less efficiently collected with funnels. Mites adapted to such naturally dry habitats may become quiescent, rather than flee, and some (e.g. Nanorchestidae) appear to tolerate very low humidities. Thus, funnel extractions from these habitats may be both inefficient and taxonomically biased.
Usually, only the larger, more active animals come out of a sample during the first day or two of extraction. Then for a few days, as the samples dries out, few or no animals are collected. As the critical level of dryness is reached, large numbers of mites will begin appearing in vials. The length of time needed to extract most of the mites in a sample will depend on the size of the sample, how moist the sample is, and the weather. For example, a 500 cm3 sample of rainforest litter should remain in a funnel for 4-5 days in dry weather, but may require a week or more in the rainy season.
Mites can be collected directly into a preservative (70-80% ethanol with 1-3% glycerine to keep the specimens moist if the alcohol evaporates) or collected live into a vial with a floor of moistened charcoal-plaster (see below). Although a good preservative, alcohol is not ideal: the longer a specimen is in alcohol, the harder and more difficult to clear it becomes. Also, volatile liquids like alcohol may deter mites from entering the funnel or pickle them in the sample. Alternatively, polyethylene glycol or glycerine-glacial acetic acid solutions may be used, but carcinogenic and potentially explosive material such as picric acid should be avoided.
Unfortunately, berleseates (i.e. the preserved samples, also called tullgrenates) often contain large amounts of soil and detritus that is knocked into the funnels by the movements of larger animals or by clumsy researchers. A layer of wide-meshed cloth (muslin or cheese cloth) or fine screening under the sample will help reduce the fall. Also baffles or other ·autosegregators· (e.g. Aoki 1984) that capture fallen soil under the sample can be useful. Dirty berleseates can be cleaned up using one of the flotation techniques described below. In very sandy soils or dry clays, samples will invariably be filled with soil particles that are mite-like in size and color, and a flotation technique should be considered.
(b) Flotation. Mites can be removed from samples of soil by taking advantage of two characteristics of arthropod bodies: 1) they have a different specific gravity than soil particles and 2) most arthropod cuticles have a strong affinity for petroleum derivatives. Specific Gravity Flotation releases mites from soil or traps when the sample is mixed into a concentrated solution of sugar (cheap, but messy) or salt. Mites will remain in suspension, while soil particles settle out. The suspension can then be passed through a filter.
Hydrocarbon Flotation is a less messy technique that takes advantage of the tendency for arthropod cuticle to adhere to liquid hydrocarbons (e.g. heptane or kerosene). A real advantage of hydrocarbon flotation is that soil samples can be preserved in ethanol and extracted at any convenient time. This also allows plant matter time to become saturated and sink. For extraction, samples are poured into a tapered, stoppered flask and a layer of an appropriate hydrocarbon, e.g. kerosene, is added. A magnetic stirrer can be used to agitate the sample to bring the mite bodies into contact with the kerosene layer; or the flask can be gently shaken to mix the kerosene through the substrate. Avoid shaking too vigorously or a milky emulsion will result. After sufficient agitation, the flask is set aside and the droplets of kerosene eventually rise to the top of the flask, bringing the mites along with them. The kerosene layer is then decanted through a fine-meshed sieve (< 200 mm) and the mites washed clean with 95-100% ethanol. This procedure should be repeated 2-3 times per sample until mites cease to appear in the kerosene layer. Detailed instructions for a more quantitative approach can be found in Walter et al. (1987). The main drawback to hydrocarbon flotation is that heavy mites, those with encrusted soil particles, and some taxa (e.g. Histiostomatidae, Nothridae) are not efficiently extracted. Also, hydrocarbon floatation requires adequate ventilation · usually a fume hood is needed.
Kethley Agitation can be used to collect mites from mineral soil or beach sand (Kethley 1991). You need a large bucket (30 L), a wooden paddle for stirring, a scoop, a fine-meshed sieve, a funnel, large collecting vials, ethanol in a squirt bottle, and lots of water (fresh or salt). The bucket is filled with water and a few litres of sand are gently added as one stirs the water into a easy but constant swirl. As the sand moves through the agitated liquid, lipids and other materials present in the soil form soapy bubbles that trap small mite bodies. The resulting foam collects on the surface of the bucket and can be scooped out into the sieve. The squirt bottle can then be used to cut the foam and move the mite bodies through the funnel and into the collecting vial. Because a large amount of sand and organic material is also collected with the mites, these samples can benefit from a kerosene flotation.
(c) Traps. Pitfall traps can be used in the field to collect some of the larger surface-active mites, and are especially effective for Erythraeoidea, Anystidae, large Oribatida, and mites phoretic on insects. Similarly, malaise traps, yellow pan traps, and other techniques for collecting flying insects will also collect their phoretic mites. Pitfalls can be baited with carrion, rotting fruit, or dung to attract particular types of insects and their phoretic mites.
Fixing, Preserving and Preparing Specimens
(a) Fixing and Preserving: Typically, mites are killed, fixed and preserved in 70-80% ethanol. In higher alcohol concentrations mites become brittle; at lower concentrations bacterial and fungal degradation can occur. However, when preserving specimens for DNA analysis, 95-100% ethanol should be used. Good seals on containers are a necessity, especially in warmer areas, and refrigeration can be used to reduce fluid loss and inhibit fungal growth. Unfortunately, as time passes, alcohol-preserved specimens tend to become tough and difficult to clear. An additional problem is that soft-bodied mites often become distorted, and the legs of many mites curl under their body when they die in alcohol. Clearing and use of gentle heating during mounting, or killing live mites in boiling water, sometimes circumvents these problems. Isopropyl alcohol (·rubbing alcohol·) or methylated spirits may be easier to obtain than ethanol, and are acceptable for killing and preservation, at least for short periods, but specimens tend to deform as they do in ethanol. Lactic acid (65%) is used by some oribatid workers, although loss of cuticular mass (especially from setae) may occur over time.
Various combinations of ethanol, methanol and glacial acetic acid have been used as preservatives. Saito and Osakabe (1992) tested a number of these formulations on mites from the families Tetranychidae and Phytoseiidae. They found that MA80, a 2:2:1 mixture of methanol (99.5%), glacial acetic acid (99.3%), and distilled water, fixed specimens with legs extended and the cuticle in good shape. Specimens stored in MA80 make good light microscope slides when mounted in Hoyer·s Medium (see below), but specimens destined for scanning electron microscopy (SEM) should be transferred to ethanol after 20 minutes to avoid deformation (Saito, pers. comm.).
Glycerine (1-3%) can be added to ethanol fixatives to protect specimens if evaporation occurs. Long term storage of oribatid mites is often in lactic acid. Water mites (Prostigmata: Hydracarina) are usually preserved in Koenike·s fluid, a smelly, but excellent all around preservative (recipe below). Rinse alcohol-preserved specimens in water before transferring them to Koenike·s and visa versa. Always remember to put data labels (printed in pencil or in indelible ink) in all collections.
Koenike·s Fluid MA80
Methanol --- 40 ml
Glacial acetic acid 10 ml 40 ml
Distilled water 40 ml 20 ml
Glycerine 50 ml ---
(b) Preparation for Light Microscopy: Identification of mites requires high magnification. Optical microscopic observation at 100-1000 x is usually needed for identification, and often high-dry (>40 x) or oil immersion objectives must be used. Therefore, proper clearing and slide-mounting of mites is a necessary skill in acarology. Some or all of the following steps may be needed: clearing, dissection, rinsing, dehydration, mounting, drying, ringing, and labelling. For specialised approaches such as scanning or transmission microscopy, a recent review of techniques should be consulted.
Clearing. A number of clearing agents are used in the preparation of mites for microscopic observation. The objective is to macerate and dissolve internal tissues, especially muscles; to soften cuticle for dissection; and increase the transparency of mite bodies. The length of time to leave a mite in a clearing agent is a function of the agent and its concentration, how long the mite has been preserved, the temperature used, the size of the mite, and the type of mite. The longer a mite has been in alcohol, the longer it takes to macerate internal tissues. Blood-filled parasites should have the body wall punctured and blood gently expressed before clearing. In contrast, freshly collected specimens may need little or no maceration, especially if soft-bodied and unpigmented. Gentle warming will accelerate the rate of clearing, while dilution tends to allow for extended periods in the agent (e.g. to accommodate a busy schedule or a weekend). One must be careful to avoid over-clearing, as this results in specimens that are brittle and likely to fall apart when handled.
Lactic acid (60-95% aqueous solution) is the least caustic and best all around clearing agent - relatively gentle and slow acting. Lactic acid is also good for temporary slide mounts and for the depression slide mounts or Grandjean carbon-block methods used to observe Oribatida (Grandjean 1949, Evans 1992). Lactophenol, commonly recommended in older texts, should be avoided because of the carcinogenic properties of phenol. For tougher clearing jobs Nesbitt·s Fluid (especially useful for old alcohol-preserved specimens) or 5-10% potassium hydroxide can be used. Both of these substances are caustic and must be used with care, both to protect oneself (especially the eyes) and to protect the mite specimens. Gentle heating (45·C) in an oven, on a hot plate, or in a water bath will accelerate clearing.
Nesbitt·s Fluid
Chloral hydrate 40 g
Concentrated HCl 2.5 ml
Distilled water 25 ml
Mounting. Three general classes of mounting media are used in acarology: water-miscible mixtures, resin-based ·permanent· mounts, and polyvinyl alcohol solutions. Water-miscible media based around gum arabic (and often glycerine) are not permanent, but have a long and uninspiring history of misuse, misformulation, and miserable results (Upton 1993). In spite of numerous problems, these essentially temporary media are preferred because of their ease of use (usually only one transfer, from clearing agent to medium, is required) and excellent optical properties. Resin-based mounts usually have poorer refractive indices and require extensive preparation, several transfers of the specimen, and often noxious or expensive solvents. Some, such as Canada balsam (refractive index 1.535), are not miscible in alcohol or water and require extensive preparation of the specimen. Although balsam mounts are supposed to be permanent, many will turn red, brown, or black over time. Some polyvinyl alcohol preparations also have a bad reputation, but more recent formulations show promise and are discussed below.
Because mounting media are often unreliable, digital-image capture is recommended as a back-up of type material in case specimens deteriorate over time. Digital-images can be considerably enhanced by layering and filtering of focal plane series of slide-mounted mites. Additionally, the use of digital images as voucher specimens and their use in expert systems, electronic identification tools, and for rapid exchange of information make them especially important in modern ecological research.
Gum arabic media. Optically, the best mounting medium for mites is a recipe usually called Hoyer·s Medium, although the proportions of gum arabic (gum acacia) and chloral hydrate vary among recipes. Chloral hydrate (also called ·knockout drops· or ·Mickey Finns· in crime novels) is a substance that may require special permits to possess and should be treated with caution. Don·t eat Hoyer·s or rub it on your skin. Crystalline gum arabic (i.e. amorphic, ·rocks·, ·tears·) may be difficult to find, but reputedly, if commercial preparations of powdered gum arabic (gum acacia) from oriental grocers or artist shops are wetted up with ethanol, they will then dissolve in water. Since Hoyer·s is hygroscopic, once the slide is dried, the edge of the coverslip should be sealed with a waterproof material (see below).
Although Hoyer·s Medium is easy to use, optically excellent, and has been the preferred choice of many acarologists, often it also is roundly cursed, because of the tendency of preparations to spoil. Most commonly, air bubbles invaginate under the coverslip as the slide dries, the medium becomes filled with oily globules, or it takes on an opaque granular consistency. In the first case, too little (or too dilute) medium was used to make the slide. After rehydration (e.g. on a wet paper towel in a petri dish), the specimen can be transferred to a new slide. The sources of the oil globules and granulations are not known, but presumably represent an interaction between the medium and some contaminant in the mite, clearing agent, slide, or sealant. Since Hoyer·s is hygroscopic it must be ringed with a varnish or paint that prevents water from entering or leaving the mount. Insulating paints are more effective vapour barriers than more readily available materials such as fingernail polish (which tends to crack). A well made and ringed Hoyer·s slide may last for decades - or it may spoil in a few months.
To make Hoyer·s, dissolve gum arabic in distilled water and let it sit over night. The next day, add the chloral hydrate and glycerine and then either filter the mixture through clean glass wool, or more efficiently, centrifuge the mixture. Store Hoyer·s in a brown bottle to prevent degradation by light; however, avoid ground glass seals and screw tops, as they can become solidly fused.
Hoyer·s Medium Heinze-PVA
(Krantz 1978) (Evans 1992)
Polyvinyl alcohol --- 10 g
Chloral hydrate 200 g 100 g
Crystalline gum arabic 30 g ---
Glycerol 20 ml 10 ml
Distilled water 50 ml 60 ml
85-92% Lactic acid --- 35 ml
Polyvinyl alcohol (PVA) preparations have been recommended by several acarologists over the years. Although some of the early PVA formulations have been shown to spoil, Heinze PVA may be a more than adequate substitute for Hoyer·s. Trials undertaken by the authors have had good results as long as enough medium is added to the slide to avoid invaginations under the coverslip on drying. Specimens can be recovered from PVA-slides after soaking in warm glycerine, lactophenol, or dilute Nesbitt·s solution (this is a good way to recycle residual clearing agent). Polyvinyl alcohol comes in a variety of preparations, some of which may yield different results. Heinze PVA (polyvinyl alcohol of M.W. 22000, Bdh: 305735b; Gohsensol GSS-5407; hydrolysis degree 30-36 mols) is mixed thus:
1. add water to PVA powder, stirring constantly in a water bath at just below boiling
2. add lactic acid and stir for a few minutes
3. add glycerol and stir until smooth
4. cool until luke-warm
5. dissolve in chloral hydrate
6. stir thoroughly, filter through glass filter paper in a suction funnel (or centrifuge)
7. store in a brown bottle
Resin-based mountants. To avoid the problem with spoilage, especially for type material, a longer lasting medium such as Euparal (eucalyptus oil and paraldehyde) or Canada balsam may be used. Specimens preserved in these media must be thoroughly macerated to remove muscle tissue and internal organs, and usually require an elaborate series of dehydrations (less so for Euparal, which will tolerate some water) and replacements, and may benefit from staining with agents such as acid fuchsine. Commercial preparations for thinning Canada balsam and Euparal are available. Canada balsam can also be thinned with xylene (under a fume hood only). Euparal (refractive index 1.483) is alcohol miscible and can be thinned with absolute isopropyl alcohol or absolute ethanol (Upton 1991). Euparal appears to be preferred by dipterologists (Halliday 1994), has better optical properties than Canada balsam, and does not turn brown over time. However, soft-bodied specimens may become deformed and distorted as the medium sets, or the preparation may craze, become powdery, or polymerize. Both Euparal and Canada balsam have pleasant fragrances. Some synthetic resins such as dimethyl hydantoin formaldehyde (DMHF, DMHFR) have excellent refractive indices and do not require dehydration of specimens (Steedman 1958, Bameul 1990), but data on how long these preparations last are unavailable.
Making a slide.
To mount a mite in medium, first place a drop at the centre of a glass microscope slide. Centering is made easier if you draw the outline of a slide on a sheet of stiff paper and place an X in the center. The drop of medium should be large enough to fill the space under the cover slip when the medium is dried. This will depend on the size of the coverslip, the thickness of the mite, and the viscosity of the medium. Hoyer·s is hygroscopic and will gain and lose water depending on the relative humidity. When drying, both Hoyer·s and PVA will contract slightly as the water is driven off. If too little media is used, then air bubbles develop under the cover slip. Transferring the right-sized drop of medium takes practice, but if in doubt, too much is better than too little. Once a slide has dried, excess media can be scraped and wiped off.
Mites can be mounted directly from water, alcohol or Nesbitt·s into Hoyer·s or PVA, but specimens cleared in KOH or lactic acid must first be rinsed in water. Mounting directly from an alkaline agent will cause a precipitation and degrading of the medium. Mounting directly from lactic acid eventually results in the growth of acid crystals within the mount. These crystals are pretty, but obscure the specimen. Also, the presence of residual clearing agents can result in degradation of the Hoyer·s over time.
Before mounting, you must decide if dissection is required to expose structures necessary for identification. Fine watchmaker·s forceps and insect minuten pins mounted in wood skewers or match sticks are the most useful tools (see Norton & Saunders 1985 on making tools). Large mites usually require dissection, and in some groups they are always dissected before mounting. Large mites can be mounted in depression slides or surrounded by protective plastic rings (Foulkes 1983), but much of the animal will not be available for observation. Also, the distance between a slide and an oil-immersion objective is limited. If oil immersion observation is required, then a slide cannot be very thick. Coverslips should be circular, appropriately sized for the specimen (10-18 mm in diameter, except for very large mites), and thin (# 000 - 1). The small (10-12 mm), thin (000-00) coverslips can be very expensive and provide less protection for the specimen, but have the advantage that a smaller area need be searched to find the mite and an oil immersion objective is easily used.
In most groups of Mesostigmata, a good lateral view of the chelicerae is usually required for species identification, and in many Phytoseiidae and Ascidae the female sperm access system is diagnostic. Crush-mounting (using tweezers to gently compress a mite between coverslip and slide) is a quick and dirty alternative to dissection, although the results often are less than aesthetically pleasing. With a well cleared specimen (and some practice), it is possible to use crush-mounting to shear away the dorsal shield from the ventral shields, expose the sperm access system, and flatten the chelicerae in a few seconds.
If the primary purpose of the slide is taxonomic research (or if you expect a taxonomist to look at your specimen) then the rule is one mite per slide. For ecological studies, however, one often mounts dozens of mites on a slide to save time and money. This can make later identification difficult if there are many taxa on a single slide. A reasonable compromise is to sort specimens into what you hope are similar groups, single-slide the mites until you have a good idea of the species present, and then go to more densely packed slides if required.
A small, stainless steel pin with a flattened tip and mounted in a wooden handle makes a good tool for transferring mites. Once the mite has been transferred from the macerating agent and through any rinsing and dehydration procedures needed, and a clean glass slide (75 x 25 mm) with a drop of medium in the center is ready, follow these steps:
1. Place the mite in the centre of the drop of medium and gently push it down to the surface of the glass slide (friction will reduce mite-movement when the cover slip is applied).
2. Decide which surface of the mite is most important for identification (this will vary with the type of mite) and orient that surface upward.
3. Orient the mite with its anterior end facing the bottom of the slide, because most microscopes reverse the image.
4. Allow any air bubbles trapped in the medium to rise to the surface and burst.
5. Holding the coverslip with a pair of tweezers, touch one end to the slide and to the meniscus of the medium. Gently lower the coverslip and avoid trapping any air bubbles.
6. If the mite begins to move, gentle pressure on the coverslip with the tweezers can be used to control its movement and orientation.
7. If the legs or mouthparts of an undissected specimen are curled out of sight, then they can sometimes be extended by pressure or by gentle heating over an alcohol flame or a hot plate (45-65·C). Wear safety glasses, don·t boil the medium, and POINT THE TOP OF THE SLIDE AT A WALL or other safe area. Coverslips can explode off slides and do injury to yourself or others.
8. Pick up the slide, invert it in your hand without touching the coverslip to any surface, and circle the position of the specimen with a fine-pointed indelible marker. This is a bit tricky, but it will save you a lot of search time under the microscope.
9. Revert the slide to its upright position, label it with an indelible marker.
10. Place the slide in an oven at or slide warmer at 45-50·C until the medium around the rim is dry to the touch. When the slide is dry, replace the temporary labelling with a permanent label with all the necessary data.
Ringing, Labelling, & Storing Slides. Once a Hoyer·s mount is dried, the coverslip must be ringed with a material that will prevent absorption of water vapour from the air. Commercial preparations of paints used to coat electrical connections (e.g. Red Glypt Insulating Varnish·, Isonel·) work well (Travis 1968), and nail polish will do in a pinch (although it tends to crack with age). Allow the slide to come to room temperature (to prevent wrinkling of the ringing compound as the slide cools) and then place the slide on a ringing stand, give it a whirl, and paint over the edge of the coverslip and a swath of the slide with a small brush or a bottle applicator (Wu 1986). Remember that the goal is to seal off any opening between the medium and the atmosphere, not to cover up the specimen. Clean the brush in the recommended solvent. PVA slides do not need to be sealed, but ringing may help keep the mount from falling off the slide.
Labelling Slides. Unlabelled or poorly labelled slides are worthless for scientific research. Make sure that you put all relevant data on the slide. Additional information can be stored in notebooks or databases and referenced by an accession number or bar code. Labels vary in quality, and some spoil in only a few months: be sure of the quality of your labels. Information can be printed in India ink or professionally printed. Laser printed labels are commonly used, but their lifespan is unknown.
When labelling, the more important information tends to be put on the reader·s left, but what is considered important varies with discipline, e.g. systematists tend to put taxonomic information on the left-hand label but ecologists tend to put collection information on the left of the coverslip. Whichever you choose, one label should have all the important collection data including the locality (preferably with coordinates), the date collected, the habitat and the collector. The other label should contain the identification information, who made the identification and the mounting medium. Use pencil for uncertain identifications. Store slides horizontally with the coverslip-side facing up, in slide boxes or flat trays.
Electron Microscopy: The study of mites is enormously enhanced by good electron microscopy. Anyone with access to a sputter coater and a scanning electron microscope (SEM) can acquire images of many mites with a minimum of preparation time and expense. Initial killing of mites should follow the guide-lines reviewed above (Saito & Osakabe 1992). Additionally, mites can be boiled alive in a test-tube of lightly soaped water and boiling chips to extend limbs, clean them of some of debris, and fix tissues. Remember to point the test tube at a wall or splatter guard in case it boils over. Traditionally, mite specimens have then been fixed in gluteraldehyde (and often osmium tetroxide) and dehydrated before critical point drying, but this approach provides numerous opportunities to lose or damage specimens and results are often less than satisfactory (Achor et al. 2001). Additionally, these elaborate techniques are not necessary for many mite taxa.
The general rule is that the better sclerotized the mite, the less treatment is required before SEM. For example, most brachypyline and ptychoid oribatid mites can be dehydrated in 100% ethanol, transferred to a small watchglass of acetone, and dried under a fume hood in a few hours with little or no resulting deformation. Specimens are then transferred to a SEM stub covered with double-sided sticky tape using a fine brush and sputter-coated with gold, gold-palladium, platinum, or carbon.
An equally simple technique, developed for preparation of internal tissue (Nation 1983), is to transfer specimens from 100% ethanol into a solution of hexamethyldisilazane (HMDS). This solvent is highly toxic, so preparations must be carried out under a fume hood using protective gear such as rubber gloves. One may transfer mites under a microscope set up under the fume hood and use a microspatula to move the specimens into an excavated block with a few ml of HMDS. When processing large numbers of specimens, a small bore (< 1 mm) pipette can be used to transfer the mites to an empty block and then to remove excess ethanol before adding HMDS. The block should then be covered with a glass plate and the specimens allowed to sit for 2-4 h (depending on the size of the mites), before removing most of the HDMS with a pipette, and adding a few ml of fresh solution. The block can be covered again for an hour, before removing the lid and allowing the HDMS to evaporate under the fume hood overnight. This technique usually produces excellent specimens of lightly sclerotized mites for use in traditional ambient temperature SEM, but soft-bodied mites often collapse or deform.
The best approach for soft-bodied mites, and especially for mites on plant tissues, appears to be low temperature electron microscopy (Achor et al. 2001). This technique has been used to produce striking images of Pyemotidae, Eriophyoidea, Tydeidae, Acaridae, and other soft-bodied acarines that are rarely, if ever, free from deformities when prepared using other techniques. An exception is the environmental scanning electron microscopes that allow the visualization of live specimens on leaf surfaces or stuck to stubs under low vacuum. Chilling the stage of an environmental SEM reduces movement and lessens the chance that the specimen will suddenly leap off of the stub.
Molecular Analysis: DNA can often be recovered from specimens stored in 70% ethanol, at least over the short-term, but killing and storage in 100% produces more reliable results. Even better is flash freezing live mites in small plastic tubes or in a buffer solution at ·70·C or lower. Standard Chelex DNA extraction technique (Hillis et al., 1996) work well with mites of moderate size using 1-5 specimens ground up with a micro-mortar and pestle. A variety of kits are on the market that digest DNA from specimens and allow the retrieval and sliding of the exoskeletons for use as voucher specimens.
Working with Live Mites
Mites have a high surface/volume ratio, desiccate easily, and must be maintained at a relative humidity similar to what they normally experience. For soil-litter mites, an atmosphere near saturation (>95% R.H.) is usually required, but mites adapted to drier habitats often need lower humidities. Wetted strips of paper towelling can be placed in a collection vial to obtain live mites from a funnel extraction, but many will be difficult to capture and remove from the towelling. A better method is to fill the bottom 20-40% of the collection vial with a mixture of powdered activated charcoal and Plaster-of-Paris.
Plaster sets as a crystalline matrix with pores large enough to admit water (and hence maintain a high humidity), but too small to admit mites. Plaster by itself is bright white and as well as obscuring light-colored mites will reflect light and cause considerable eye strain when observing mites under the dissecting microscope. However, a mixture of 1 part powdered activated charcoal: 2 parts plaster (by volume) will set to a uniform light grey colour that is easy on the eyes, and may also help to absorb toxins (the original reason charcoal was used in this mixture). Use only enough water to make a thick paste the consistency of cake batter. Too much water results in a loose and crumbly plaster; too little water results in rock-hard plaster that will not absorb much water. Powdered charcoal is a messy material, so a lab coat and latex gloves are useful. Other things to consider include the working distance of your microscope: vials must be small enough to fit under the microscope objective and need to be straight-sided (i.e. without necks). Shell vials and urine specimen containers work well. Modified Roberston cells drilled from perspex or other material are another method for rearing and observing mites (e.g. Solomon & Cunnington 1964).
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