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Lab & Field Techiques for the Study of Spiders

 

Contents

ATOL Default Preparations
Dissection and Mounting Tips & Gadgets
Field Gadgets

New items
Stabilizer for dissections

 
ATOL Default Preparations See preparations page in
ATOL Standard Views Manual

1. Female in alcohol

Make first the images of the entire specimen (e.g., habitus with legs). Remove left legs and palp, make lateral view without legs and the legs themselves. Reserve left palp and legs I and IV for SEM. Remove right palp and legs, abdomen, and chelicerae. See details below for removing the abdomen, carapace, legs and palp.

2. Carapace, endites, labium (female)

Remove the abdomen by cutting around the pedicel, leaving pedicel sclerites connected to the carapace. Straight any dent that may have occurred during dissection of chelicerae; often the dent will simply spring out when pressing the carapace from the sides.

If the tapeta and dark eye cups remained below the eyes (sometimes they come together with the chelicerae), this is a good moment to clarify and examine the tapeta (see preparing tapeta).

Clean the dorsal side of the endites and labrum, as there often accumulates dirt and vomit. Send to CPD. Once dried, bend or tear the remaining cuticle of the abdomen to expose pedicel sclerites, brush and mount on adhesive copper tape. Attach from a coxa, secure with conductive paint, and position.


3. Chelicera (female)

One of the most difficult dissections is removing the chelicerae without denting the carapace during manipulation, especially in hardened specimens, or to damage the chelicerae themselves. Examine the insertion areas by pushing the chelicerae slightly apart from the endites, and from each other. With the micro-chisel tool, cut the surrounding cuticle and the muscles below; leave the chillum with the carapace. Insert the chisel between basal condylus and clypeus, going deep to cut the powerful muscles. Push the chelicerae from several sides (e.g., the posterior side, or posterior-lateral insertion), to remove them. Take care not to damage the fang, teeth, and retromarginal setae. Both chelicerae will come together, firmly united at the articulation, especially at the postchillum. If the venom gland comes attached, keep them united to the chelicerae. Cut the articulation between chelicerae, leaving the postchilum on the left side; use left chelicerae for mounting, reserve the right in a micro vial with the specimen.

Remove excess of muscles, and expose the venom gland if it came attached. If the fang has mobility, open it to expose the fang furrow; push with fine forceps from the middle, not from the tip; do not force it too much, because the tip breaks easily. If the articulation is so flexible that the fang returns to its position, push the fang open after CPD.

Send the left chelicerae to CPD. Clean, and mount on adhesive copper tape, attaching from the severed base. Secure with conductive paint, and position.

4. Palp (female)

Check the position and shape of the claw, depilate the retrolateral side to expose the entire claw.

After CPD, spread it open if necessary, clean and mount on adhesive copper tape attaching from the trochanter base. Secure with conductive paint, and position. Double check that there are no loose hairs or debris stuck to the claw.

5. Leg I, female

Remove the leg from carapace by cutting between trochanter and coxa. It is easier to puncture the posterior side of the articulation with the forceps, and push the leg apart from there.

Remove the hairs from the retrolateral side to fully expose the claws; remove as well some of the long dorsal hairs if necessary. Try not to remove accessory claws, or any other informative structure beyond necessity. If there are claw tufts or tenent setae, remove the retrolateral ones to expose insertions. Take the setae off a small group at a time; otherwise the pad may tear off.

After CPD, clean, and extend the leg (remember that we will take a ventral view of the tarsus (SV129), and mount on adhesive copper tape attaching from the trochanter base. Secure with conductive paint, and position. Double check that there are no loose hairs or debris stuck to the claws or the tarsal organ area.


6. Leg IV, female

Similar as with leg I.

7. Epigyne and vulva trypsine digested

Depilate and take apart copulatory plugs. If the plugs are too strongly attached, try again after digestion. Cut the soft cuticle around the epigyne with micro knife. If the specimens are very soft, the micro-chisel tool may help to pierce the cuticle. Be sure to cut the cuticle until the epigastric fold (but remember that some groups have structures posterior to the epigastric fold that we may want to image as well; see preparation 9). Remove the epigyne, usually pushing from one of the sides, close to the epigastric fold. Double check that it comes apart clean, because it is easy to tear off a long stripe of cuticle from the epigastric fold. Finish the shaving, and send to pancreatin digestion. After digestion, clean loose tissue remains with brush, jet cleaner, and ultrasound cleaner. This is a good moment for clarification and illustration with compound microscope.

After CPD, remove or bend any soft cuticle hiding the internal genitalia (e.g., a booklung atrium). Sometimes it is necessary to break part of the posterior margin of the epigyne to fully expose the spermathecae (Fig. ...). In that case, break only one side. Clean and mount on adhesive copper tape attaching from the soft cuticle at the side or before the epigyne. Secure with conductive paint, and position. Double check that there are no loose hairs or debris stuck to the claws or the tarsal organ area.

8. Spinnerets female

This is one of the more informative preparations, so it pays well to be careful. The most important step is selected a clean specimen, with the spinning fields well preserved, not collapsed and preferably well exposed. This may be contradictory, since exposed spinning fields may wear-off by friction with the vial and labels. If the sorrounding hairs are lost, especially on the PLS, this may indicate that the spigots are damaged as well.

To properly expose the spinning fields the spinnerets should spread out, and some hairs should be removed. Depending on the hardness of the abdomen, the spinnerets may remain spread out after pressing the sides of the abdomen with forceps, but this is more the exception than the rule. Anyways, we can position them again after CPD.

The difficult part is removing the hairs. If they are too close to the spigots, leave them and depilate some more after CPD, where we can bend the hair apart from the spigots and it will not come back to its original position; the single hair tool is useful for this. If you are unfamiliar with shaving spinnerets, start first with the larger and more common species, as it requires experience to tell apart hairs from spigots. Don’t even try to do so without the finest forceps. While shaving, clean often with jet cleaner, and brush.

Shaving can be done before and after removing the spinnerets from the abdomen. To remove the spinnerets, section them by the surrounding cuticle. If the tracheal spiracle is close to the spinnerets, include it in the preparation.

After CPD, and several cycles of shaving and cleaning, mount the piece on an adhesive carbon tab affixed to a stub. Spread the spinnerets apart again if necessary, using the glue of the tab and pressing here and there. If you touch the piece, brush again to remove loose hairs, and remove any other debris with the air jet and single hair tool. Once in position add conductive paint (Fig. ...).

Sometimes it is not possible to depilate or expose the spinnerets as they come, and it is necessary to cut the group longitudinally. In that case, after dissection from the abdomen, section with the micro knife, leaving e.g., the left spinnerets, entire anal tubercle and entire cribellum on one side, and the right spinnerets on the other. Then mount as above.


9. Abdomen digested

This preparation will show the internal respiratory system, as well as any other cuticular structures, as female genitalia, uterus externus, apodemes, and the distal part of the spinning gland ducts. The soft tissues are removed with KOH digestion or pancreatine digestion. If you are unfamiliar with the technique, try some common spiders first, as it takes some tries to get used to it. If the main objective is the respiratory system, then immatures or males can be used as well; so far there are not known maturity or sexual differences in the respiratory system beyond early spiderlings. Tracheae are best examined in compound microscope, so you may want to depilate the area below the tracheal atrium.

Remove the abdomen, and make an incision to take the roof of the abdomen. In some groups the tracheae run close to the ventral surface, but in others they bent over the dorsum; some may have interesting genitalic structures extending considerably to the dorsum. That means that if you don’t know what to expect inside, cut only superficially. Sometimes the dorsal cuticle comes easily apart; it will get loose after the digestion anyways. Also, you may be interested to follow the tracheae to the carapace, and in that case include it in the preparation.

After digestion, cleaning the preparation may be tricky, as the silk glands may remain undigested and entangle with the tracheae, or the tissues may not fully digested. It helps a lot to immerse briefly in chlorazol black, which stains the interior side of the cuticle. To the compound microscope the staining is not obtrusive, and for the stereo it looks nice.

See below for tips of dark field illumination under the stereo.

Don’t use the brush to clean the tracheae, as you will entangle and tear apart them. The jet cleaner is useful here, always in the direction of the tracheae, from spiracle to tip. It is common to find oil droplets, especially on fresh specimens; these dissolve after a day or two in alcohol.

To observe the tracheae, place the preparation on an excavated slide. I prefer lactic acid as media, because it clarifies the remaining dirt and silk glands, while the refraction index is sufficiently different from the cuticle. (Perhaps methyl salicylate works as well?)

10. Male in alcohol

Similar as with female

11. Abdomen male

This preparation can be mounted in one piece, although the complexity of exposing spinning fields often requires that the spinnerets are dissected apart; follow the procedures as with the female spinnerets. Brush the epiandrium and make sure that there not remain any dirt hiding its spigots. After CPD, clean and mount on adhesive copper tape attaching from the side of the abdomen (leave the booklung spiracle free). Secure with conductive paint, and position.

12. Palp, SEM (male)

Dissect the left palp. If it can be extended to expose the ventral side, then it is good to cut between trochanter and coxa; the micro-chisel tool may help here. Otherwise, cut between patella and femur. Sonicate, depilate to expose well the copulatory organ, brush. After CPD, clean, double check that there are loose hairs stuck in the bulb, extend the palp (remember that we will take some ventral views), and mount on adhesive copper tape. Attach from the trochanter or patella base, depending where you sectioned it. Secure with conductive paint, and position.

13. Palp, expanded (male)

Dissect the palp, sonicate, brush, but don’t depilate. Prepare two dishes, one with distilled water, the other with a weak KOH solution. Transfer the piece to the KOH, after some minutes see under the stereo. We will cycle between KOH and distilled water under fully expanded, and then leave it in distilled water. Small, fresh, and soft spiders will expand more quickly, so the cycles are shorter. Larger or harder pieces need longer cycles. At any rate, don’t forget the piece too long in KOH, as it will digest everything inside and may not expand any further. For extra-hard pieces, sometimes the heat of a lamp in the distilled water phase helps expansion.

It is tricky to draw or image the expanded palps, as they continue expanding or rotating. Leaving it overnight on distilled water may help stabilize the preparation.

 

Tapeta ...
Dissection and Mounting Tips & Gadgets
Adhesive carbon tab: ...
Adhesive copper tape: We have experience with the EMS 77802 copper tape, although there is a variety of new tapes now. One of the sides is glued and covered with a a paper to peel. Cut a piece ca. 15 mm long, tapering at one end to a thin wire. Paste the thick end to the stub; press with the back of the forceps for good adhesion and flat background. Straighten the thin end, perhaps slightly bending the thin tip. The glue remains sticky for a long time, so you can prepare one or two boxes in advance. Instead of taking the sample to the copper, it is often safer to take the stub with the forceps, and touch the piece with the glued thin end. Sometimes only a few hairs adhere and it is necessary to press the piece to glue it correctly. As the glue is not conductive, and the contact is made often through hairs only, it is much safer to add a patch of conductive paintgoing from the sample cuticle to the non-glued copper side. This also helps securing the piece. The glue loses much of its stickiness after vacuum sputter coated. This is handy, since the manipulation of the thin glued stripe may cause the sample to spring off.
Air jet: This is a glass tube stretched to make a capillary and connected to a rubber tube. Blowing produces a thin current of air and is useful in removing loose dirt in dry preparations (invented by Gustavo Hormiga, GWU).
Blunt tip tool: Used for delicate manipulations while avoiding undesired punctures. Take a stainless steel minutien pin (e.g., FST 26002-15), and round the tip with the finer sand paper. Bend in an angle and mount on a interchangeable tube. It is handy for manipulation of samples in excavated slide under a cover slip.
Brush: We have experience with thin brushes of synthetic fibers. They are inexpensive. Check under the stereo scope that the fibers have a very thin, straight end. The brush is used to clean samples in alcohol or after drying. Always brush in the direction of the setae, otherwise you risk removing or placing them in unnatural positions. The brush is also handy for manipulation of dried samples, which are hard to manipulate with forceps without denting them. Slightly divide the brush in two with the forceps, and then release while embracing the sample with the hairs.
Chlorazol black: Stains the internal side of the cuticle after a digestion. We use a vial with CB saturated in 75% alcohol. Briefly immerse the piece (held by forceps) for a few seconds, then transfer to a dish with alcohol and remove excess with the jet cleaner.
Clarification: We have experience with clove oil, lactic acid, and methyl salicylate. MR recommends clove oil for heavily sclerotyzed structures (entelegyne spermathecae, male palps) and lactic acid for tracheae in digested abdomens. GH uses …
Conductive paint: MR has experience with colloidal graphite on isopropanol base (EMS 12660), but there are others as well. Mix with extender so that it is easy to manipulate with the graphite painter pin. If it is too viscous it dries before reaching the sample; if it is too fluid it will spread over a larger surface and may ruin the preparation. The conductive paint avoids charging of the sample under the SEM, and secures the piece.
CPD sample basket: A porous recipient to store many specimen holders for a batch of CPD. It is important to use internal layers or something similar to keep all stoppers in place; they will pop-up otherwise. If you use specimen holders of different sizes, an intermediate layer of aluminum foil will support the stoppers in place. A 35 mm film container is a good diameter for the chambers of several CPD brands. You can process e.g., 15 samples in one layer, or 30 in two. Don't forget to identify each sample with small labels.


CPD sample holders: They must allow circulation of liquid, and must remain closed under the pressure variations in the CPD chamber. Small specimen holders allow for larger batches of samples. Remember to add small labels with identification codes inside each holder. I usually process 15 or more samples by CPD cycle, using a CPD sample basket. My standard holders are plastic micro vials (Bioquip 1133NC) perforated by many holes; if you want to make many of them, it makes sense to have a perforator with many pins. As a stopper, I use the larger half of a silicon micro vial stopper (for Bioquip 1133L), but now they sell stoppers specific for the 1133NC. Anyways, don’t trust those stoppers, they may pop-up inside the CPD chamber; use a secure CPD sample basket. The perforated plastic vial can be cut at the middle to make an even smaller size, which allows for larger batches. The microporous specimen capsules (e.g., EMS 70188) are nice, secure, and affordable, good for larger specimens. For extremely small samples such as embryos, I have used glass micro vials with a cotton stopper (and something to keep the stopper in place).



Critical point drying (CPD): MR recommends processing all SEM samples with CPD, even if heavily sclerotized. Besides good practice, air drying often produces that the setae bend to form clusters in unnatural positions. It seems that the CPD can be avoided by using HMDS (but mind the venting); Bernhard Huber (ZMFK) uses HMDS.
Dehydrate in alcohol: Absolute ethyl alcohol is commonly used as intermediary in CPD. It is handy to have a series of jars with increasingly concentrated alcohol, e.g., 85%, 90%, 95%, used 100%, clean 100% (label the jars, not the lids). From time to time (e.g., twice a day, or every hour), transfer the CPD sample holders from one jar to the next. They accumulate in the clean 100%, ready to go to CPD. Once a batch is completed, replace the clean 100% alcohol.
Depilation: Some hairs must be removed to expose structures. Use the fine forceps to remove hairs. A good dish and proper positioning of fingers makes a lot of difference in your pulse. The stabilizer for dissections may be a great aid for removing hairs.
Excavated slide: Used for observation in compound microscope, usually with a clearing media. We use strips cut from glass covers to retain the sample in position. The sample can be positioned behind the cover using the blunt tip tool. The closer the sample to the cover, the more magnification can be used in the microscope.
Forceps: You can tell many things of a morphologist by examining his or her forceps. MR uses a robust and a delicate one for each hand, four in total. The robust forceps are for general use with specimens and vials, and for positioning the whole specimen. The thin forceps are of high quality, and are always stored protected with caps (cut the tip of a disposable pipette to make a good cap). Compare the tips in the figure (...). In my experience, a good compromise price/quality is the Dumont 55 Biology tip (FST 11295-51, 11295-20, $40-50). Fine forceps are expensive, so we take great care of them. See the maintaining forceps section. Advice: never lend your good forceps.
Graphite painter: Made with a entomological needle with blunt, slightly curved tip.
Interchangeable system for tools: The micro tools are so delicate that their storage and transport becomes a problem. As they proliferate, it may be unpractical to have individual handles for each. I use a modular system that solves both issues. Each tool is mounted on a short plastic tube, which fits at the end of a stick. I use pieces from the plastic straws of cotton swabs, one end melted on a flame to fix the tool. The stick has a thinner end that fits the tube tightly. It is important that the stick is about as thin and long as the handle of a fine brush: it has to be thin to allow rotation on your fingers, and long to make it stable. For transport the tools can be stored in vials with a toothpick on the stopper. It is wise to draw a schema of the tool in the stopper.

Jet cleaner: Used to clean loose debris in immersed preparations. It is made of a disposable pipette stretched after heating. It takes some practice to do it well. The thin outlet has to be properly cut to produce a straight, thin jet without turbulence. Use it with the same medium as your preparation (alcohol, distilled water). After a while the bulb cracks and must be (at last) disposed.

KOH digestion: Used for removal of non-cuticular structures. Always use a double boiler. Don’t heat undissolved pellets; otherwise it is easy to blow the entire preparation. If you don’t have a hot plate, the heaters for anti-mosquito tablets are just ideal and very cheap. Place a solution of KOH in water, mixed thoroughly (otherwise the KOH will remain very concentrated at the bottom), place the piece and heat for some minutes. Peek from time to time, the largest take longer. Don’t leave the sample in KOH too long, as it will bleach, weaken, and eventually dissolve. After digestion, the sample can be stained in chlorazol black. See also pancreatin digestion as an alternative.

It is wise to digest the smaller samples (spiderlings, minute spiders) inside a glass microvial with a loose cotton stopper, otherwise it is easy to miss the specimens when they become transparent. Use a microvial fitting tightly transversal at the bottom of the containing vial with KOH solution, to retain the cotton in place. Make all the changes of fluids through the cotton, under the stereo.

Manipulation of digested pieces under the stereo is easier with a dark field illumination, especially if they are delicate: place a transparent dish with the sample in alcohol over a small mirror, and point the illuminator so that the sample gets the incident light obliquely from below. A polished black plate also works, although not as nicely.


Maintaining forceps: A good forceps has very fine tips, a secure grasp at the very tip, and they won’t open with extra pressure. From time to time it is necessary to correct the shape of the tips with sand paper, always controlling under the stereo. At that small scale, a very small pressure eats quite a lot of metal. The gold rule: always retouch the tips while closed together, so their sides are even; laterally uneven tips are difficult to correct. To smooth the internal surface of the tips, work grasping slightly both sides of the folded sand paper at once. Use a block of post-its to support the sand paper to allow for a good angle under the stereomicroscope. The tips should be convergent at first contact; if they are close to parallel, they will spread open with further pressure. Press each tip separately on a glass to slightly bend them if needed.
Micro-chisel tool: Used for frontal puncture incisions. Take a stainless steel minutien pin (e.g., FST 26002-15), and press hard with the edge of the forceps on a glass, pass more times until you get a flat, sharp edge. Mount on a interchangeable tube. The pin is so thin that it can penetrate in the softer cuticles without dents. It is also used to reach far between chelicerae.
Micro knife: Made of a razor blade. Cut pieces in diagonal with an old scissor, or break with tweezers. It is easier to use the breakable blades specific for making micro knifes, although they seem to be not so sharp as regular razor blades for shaving. The much finer blades of safety razors are excellent, but require dismounting. Anyways, after cutting or breaking, the acute tip of the tool will be somewhat bent. Give it a few passes of sand paper to remove that portion, and fix to an interchangeable tube. The angle of the sharp edge is important. Acute angles are better for most dissections, but always have some knife with the appropriate angle to cut flat on your plastic dish.
Micro vials: Glass microvials are nice. They don’t come too clean, and they may be difficult to fill in. Store them in a jar with alcohol 75%, shake vigorously to fill them all, and change the alcohol. They will remain filled and clean. Don’t knead the cotton stopper with your fingers, this contaminates everything with skin cells. Instead, have a jar with 75% alcohol and a piece of loose cotton inside. Use premium quality cotton, which produces fewer loose fibers, and manipulate with the forceps only. Too expensive? see the improvised micro vials.
Micro vials, improvised: If you live in this part of the world, there may be times when you can’t afford glass microvials. I have used blue straws from cotton swabs to make cute small microvials. Tin solders for electronics are ideal to melt the plastic and seal one end. Remove the tip of the solder to make a flat hot surface, and with some practice you can make several dozens in a couple of hours.
Needle marker for SEM stubs: Engraved codes can be read in the SEM screen, which is important if you load several samples in a same SEM session. In a 0.5 mm mechanical pencil, use a sewing needle instead of graphite. Scribble preparation codes on the aluminum SEM stubs.

Pancreatin digestion:
Plastic dish: I use blue plastic dishes for dissections (actually, lids from various sources). The blue renders the colors nicely, and is dark enough to work under potent illumination. The plastic is soft enough to hold the forceps in place, allowing a secure and stable grasp. The soft plastic also protects the tips of fine forceps. It is important that the dish has a diameter and height to hold your fingers comfortably while driving the forceps.
Positioning of samples on flexible mounts: SEMs vary in the degree to which the specimen holder can be tilted. If your SEM can be tilted 90 degres, then you may be able to take all views without need to bend reorient the preparation. Many SEMs can only tilt 45-60 degrees, and in that case you need to reorient the piece at least once to image opposite sides. It is useful to orientate the sample in such a way that all the views can be taken in two sessions, with only one reorientation in between. A typical orientation is 45-45-45 degrees, so you can take the ventral, retrolateral, and apical view in one session; for the next session, bend the wire and take the dorsal and prolateral views.
Sand paper: Sand paper is used to build and maintain forceps and small metallic tools. Keep two small sheets of sand paper in a plastic bag, folded with the abrasive outside. One of them the smoother you can get, the other slightly coarser. The smoother sand paper is finer, cheaper, and easier to use than abrasive stones.

Single hair tool: Used to remove small debris in dry preparations, bend hairs away from spigots, etc. Take one fiber from a brush, and crazy-glue to a minutien pin. Bend the pin and affix to a interchangeable tube.
Stabilizer for dissections: Ever wasted specimens cleaning them for SEM? Trembling fingers this morning? Not every day is a good day for dissections! Resting your fingers on the sides of the dish helps stabilizing the forceps, but this may not be enough for the tiniest dissections. An additional piece closer to the specimen for resting the forceps helps stabilizing a lot more precisely, at the expense of a smaller range of action. I use it only for the most delicate dissections, as tearing off one claw tuft in a small leg to expose the claws, or taking off hairs to expose spigots. It's made of the a rubber stopper for patent lip vials. See the bottom images, with a long exposition, before and after the stabilizer. (Of course there is no use to it except you have the thinner forceps; see scales in images.)




Thick pipette: A disposable pipette cut to make a wider opening. Handy to transfer small to medium-sized specimens.
Ultrasound cleaner:
Field Gadgets  
Pocket labels case: You will love these slim, recycled fancy cases for labels. Prepare a good quantity of labels (blank or printed with codes) and refill every day or two.