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Journal of The LEPIDOPTERISTs, Society
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Volume 18 1964 Number 4
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NOTES ON COLLECTING, REARING, AND PRESERVING LARVAE OF MACROLEPIDOPTERA
by Noel McFarland
Valyermo, California, U.S.A.
Introduction
The purpose of this paper is to present a general outline, along with some details, which I have found useful in collecting, rearing, and preserving the early stages of Macrolepidoptera (emphasis is on larvae). The information in this paper is drawn from the author's personal experience in southern California, northeastern Kansas, and western Oregon, unless otherwise indicated. When details are given, these often refer to certain moths, which sometimes have peculiar requirements that make them more difficult to rear than most butterflies.
If more lepidopterists obtained good results from their attempts at larval rearing and preservation, there would undoubtedly be more interest shown in life history studies and similar work. The great advantages in the presevation technique outlined here (using a solution which I call "K.A.A.D.L"), are that (1) no specific time is required in the preserving (or fixing) solution, (2) no changing of solution concentrations is necessary, and (3) the results are excellent. This technique makes collecting and preserving larvae a simple matter, even when travelling by car. However, for best results, it is necessary to follow the technique in all its details, from injection of larvae (which is a quick and easy process), to the permanent storage of specimens in the larval collection.
Collecting
Beating shrubbery or trees, or sweeping through herbs and grasses with a net, produces a great variety of larvae. It is important to do this both in the daytime and at night, since many nocturnal feeders will be completely off the foodplant during the day, or they will be too low on the stems or branches to be affected by beating. If a certain species is desired, careful searching on the correct foodplant(s) will often produce the best results; again, nocturnal searches should be made as well as
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diurnal. A large flashlight or lantern, with a large spot-beam, is the most useful source of light. Searching will give much better results than beating or sweeping, where nest-building larvae are involved. Furthermore, such larvae are usually conspicuous because of the nests they build (i.e., webs, curled or rolled leaves, several leaves drawn together with silk, etc.).
If the adult is diurnal, it is often possible to watch the female ovipositing, or at least to gain clues on where to search for the larvae at some later date. When searching for larvae that feed on a very abundant species of plant, it is often much better to search on isolated plants, or in small clumps, rather than searching in the midst of a large colony of the plant. Where trees are involved, small saplings at the edge of a forest grove, in a field, or along roadsides, will often produce excellent results. When collecting on plants that have stiff, leathery leaves (i.e., various sclerophyllous chaparral plants) it is usually necessary to search during the limited season when new growth is still soft, or when flowers are present. The larvae feeding on such plants often refuse the older (or hardened) growth. An example of this condition is seen in Quercus agrifolia (coast live oaks in southern California.); most of the numerous larvae that feed on this tree are present only during that short season (March-April) when the new leaves are growing and tender. These larvae grow rapidly, and most of them will have left the tree by the time the leaves have hardened.
Some pupae may be collected by digging in the soil near or under the foodplant, or by looking under logs, boards, rocks, etc. Other pupae may be discovered in leaf litter that accumulates in the crotches of tree trunks, or around the base of a tree. Some species spin cocoons in bark crevices, or under loose bark. If burlap bags are tied around a tree trunk, certain nocturnal larvae will hide unde these by day, and others may pupate in or under them. Larvae that feed on low-growing herbs may sometimes be attracted to boards placed on the ground, and can be collected in the daytime, when they are hiding under the boards.
Rearing
Whenever possible, it is preferable to rear a species from the egg, as this will provide representatives of all the larval instars, and will show variation within the species. It is a good idea to start with 50 or more eggs, if time and space permit.
Moths will often oviposit readily in captivity. With many, all that is needed is confinement in a box (many saturniids, some sphingids, many arctiids, and others). Many noctuids, notodontids, geometrids, and others will oviposit on strips of stiff (i.e. starched) cheesecloth, in a small glass jar. Regular cheesecloth, or strips of paper towel, may be
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satisfactory in some cases. Some will oviposit inside of brown paper bags (Catocala). Others require sprigs of the foodplant (with fresh leaves), or stems, or bark of the foodplant. These species often require more space (in order to fly around the foodplant) while ovipositing; most butterflies and diurnal moths fit this category, as do some nocturnal moths. Many confined moths will require feeding, as they may only lay a few eggs per night, and will not live long without some liquid. A mixture of one part sugar (or honey) and two parts water is satisfactory for most of those that require feeding. This should be offered to the insect at least every 48 hours, a bit of cotton is saturated with the solution, and the moth or butterfly is placed on the wet cotton-ball. If it does not unroll its tongue to drink, it should be held while its tongue is carefully unrolled with a fine insect pin; when the tongue is touched to the wet cotton, feeding will usually begin. As a more nourishing source of food, for species that must be kept alive for many days (or even weeks) in order to get oviposition, Dr. John G. Franclemont recommends the large, "sticky" Del Monte raisins, which may be soaked in the sugar-water solution. The same wet raisin may be used many times over, as a source of food. (Peculiar requirements for oviposition will sometimes be encountered. The above discussion mentions only the easiest methods, which are successful in numerous cases). Where diurnal species are involved, sunlight is often a requirement, as well as regular feeding, and the presence of fresh sprigs of the foodplant. Means must be devised to provide sunlight without killing the insect from overheat, and the plant material upon which it is to oviposit must be kept from wilting. The container used may be a jar, with cheesecloth covering the top, and a piece of thin white sheet partly shading it. Variations can be worked out to suit the species. Sometimes, small screen cages are better.
Eggs which overwinter are easily kept in good condition if they can be housed in an area with a climate similar to that of the collection site. The jar in which they were laid should be kept outdoors until the following spring, when food is available. They should be kept out of sun or rain, in a covered shed or garage, but exposed to natural outdoor temperatures. Overwintering larvae and pupae are easily handled in the same way. The eggs should not be brought inside until foodplant leaves are well-started. Very young leaves are sometimes sticky, or otherwise unsuitable for small larvae. The eggs should be kept in clean jars, where they were originally laid, and the jar lids should not have holes. Excessive dryness or any condensing moisture in the jar, should be avoided. If the eggs were laid on fresh leaves, the individual leaves with eggs on them should be separated and allowed to dry out somewhat before being
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closed up in a jar; if the jar "steams" inside, it should be opened for a short time. As larvae begin to hatch, they may be transferred from the egg-jar to another jar, containing samples of the foodplant leaves, of varying age and tenderness; the remaining unhatched eggs, which are nearly ready to hatch, should NOT be placed among green leaves in a humid jar, or they will often fail to hatch. Small jars with solid lids are best for starting larvae; in these jars, they are easily cared for, and they will not become lost. When larvae are first beginning to feed, they occasionally need to be placed on the leaves several times, or they will wander around and finally starve. A small water-color brush is useful for transferring newly-hatched larvae. If the foodplant is unknown, first try any plant eaten by a related species of moth or butterfly; also, try other plants in the same family as known foodplants. If no clues are available, the following generalization is sometimes helpful, in that it eliminates a great number of plants that one might try: moths that lay "large" eggs, for the size of the moth, nearly always feed upon some woody tree or shrub; those that lay "small" eggs usually feed upon herbaceous plants, weeds or grasses, etc. This phenomenon seems to apply in nearly every case! In offering plants, the following plants, or near relatives, are worth trying in the U.S.A., in addition to others peculiar to certain localities: (a) woody types — Quercus, Salix, Populus, Ulmus, Juglans (or other nut tree), Arctostaphylos, Fraxinus, Alnus, Rosa, Rhus, Ceanothus, Rubus, Ribes, Prunus, Crataegus, Cercocarpus, Cornus, Robinia (or other woody legume), Eriogonum, Sambucus, Lonicera, Symphoricarpos, Pinus, Juniperus (or other conifers), etc. (b) herbaceous types — Brassica (or other crucifer), Mentha (garden mint), various clovers, Lotus, Chenopodium, Polygonum, Plantago, Fragaria, Asclepias, Malva, Galium, Penstemon, Oenothera or Epilobium or Clarkia, Pteridium (or other fern), Arctium, Aster, Helianthus, thistle (or other composites), an annual grass and two or more coarse perennial grasses, etc. The above plants will not, of course, suffice in all cases, but one of them (or a close relative) may often be acceptable. When various plants are tried, only one or two leaves of each type should be offered, to make sure the larvae will be able to crawl over all of them with ease. Tender leaves should be offered, but not extremely young leaves.
As the larvae grow, they may be transferred to larger jars, screen cages, or "sleeves" of netting upon the foodplant (outdoors). At all times ample food should be available, and crowding should be avoided. If disease does not kill overcrowded larvae, the resulting adults are likely to be dwarfed. Half-pint, pint, and gallon jars (wide-mouthed) are suitable for rearing most larvae, unless very large numbers are being reared. Lids without holes should always be used. The obective is to
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keep the jar somewhat humid or "steamed" inside, which in turn keeps the foodplant leaves fresh for several days. Jars should be opened to air every day or two. When the larvae are placed on fresh foodplant, the jar should be throughly cleaned under hot water from the tap. An aluminum baking tin is a very useful container into which to dump larvae prior to changing them; if it is about two inches deep, and smooth, it is difficult for them to crawl out, and the tin is readily washed with hot water, after use. When using jars for rearing, it is necessary to pay close attention to conditions in the jar, and to air them out regularly, or disease may develop. Change the foodplant before it runs out, or when the jar becomes too dirty with frass. Rearing-jars should be kept in a well-lighted (but sunless) location, at 70°F., or less. A gallon jar is suitable for about 20 - 50 (depending on size) average noctuid larvae in last instar. In most cases, the jar-technique is convenient and time-saving, and the larvae usually grow rapidly.
Some larvae definitely require sunlight and/or ventilation. Examples of larvae with these requirements are certain arctiids, among which are Apantesis, Arctia, Haploa, Kodiosoma, and Platyprepia; many saturniids (Hemileuca., Pseudohazis, Calosaturnia, and others); many sphingids (after third instar); some lasiocampids; a few geometrids (particularly larger types, such as Bistort and Cochisea); most papilionids; some nymphalids (Euphydryas and Chlosyne), etc. Such larvae are best kept in screen cages or other ventilated containers. Those that also need sunlight will feed and grow well if this is provided for at least two hours daily. (If necessary, electric lights may be used in place of sunlight. ) The foodplant is kept fresh in a small jar or tube of water, which should be plugged to keep larvae from crawling in and drowning. The "sleeve" technique, outdoors on the foodplant, is especially useful when rearing large numbers of one species. Tough nylon or dacron netting makes a good outdoor bag, for use as a sleeve. White bed sheets can also be used; they keep birds from seeing the larvae, and give protection from too much sun or wind.
In handling larvae, those that cling with great tenacity to the plant stems( most sphingids, saturniids, and some geometrids, etc.) should never be forcefully pulled off, or the prolegs will be injured; such larvae may then bleed to death. Any larva that is ready to moult should be left where it is; if dislodged from its silken mat, it may be unable to pull free from its old skin, and will die when moulting is attempted. Larvae about to moult are very easy to recognize because of the swelling of the new head capsule under and behind the smaller old one-such larvae will remain in exactly the same place for two or more days.
Pupation requirements of larvae vary greatly. Among the moths, a
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large number of species (especially sphingids, noctuids, notodontids, and some geometrids) require soil, into which they burrow. This must be provided when the larvae stop feeding, and begin to crawl around the bottom of the jar. Three or four inches of damp loam, in a gallon jar, is suitable for 20 or 30 "average" noctuid larvae. A layer of leaf litter, about one inch deep, should be provided on top. The soil should be damp enough that it will not cave in when the larvae burrow into it, but it should not be wet. After about 14 days, the pupae should be dumped out, and stored in special containers for pupae, where they will be less exposed to attack by fungi. Other larvae have a very special requirement of soft wood or pith, into which they chew for pupation. Such larvae will die without pupating, if placed on damp soil. (Examples are agaristids and noctuids of certain genera, such as Alypia, Psychomorpha, Raphia, Behrensia, Fleroma; notodontids such as Centra). A good material for such larvae is yucca stalk pith, split lengthwise. Also useful are pieces of Celotex, or similar material, or strips of fibrous, stringy bark. Numerous geometrid larvae spin slight cocoons within loosely-curled leaves, either on the ground or on the foodplant. Most saturniids and arctiids, and a few noctuids and others, spin cocoons above ground; such cocoons offer no special problems. The same may be said of most butterfly chrysalids. Naked, underground pupae, that do not emerge for many months (i.e., aestivate and/or overwinter), present problems in keeping them alive during this long period. If kept too damp, they are often killed by fungus, or they rot; if kept too dry, they dry up and die. In general, it is safer to tend toward too dry than too wet conditions; during most of the diapause period, near-dryness in a closed container produces good results. Pupae should not be kept in a heated room, or where the air is very dry. When it is time to break the diapause, warmer temperatures and damper soil are usually needed. In general, the safest way to handle over-wintering pupae is to leave them outdoors most of the winter, if they are native to the area, or to another area that has colder (or equally cold) winters; if early emergence is desired, they can be brought inside in late winter, instead of waiting for normal warming outside. Uniform cold, as in a refrigerator, gives very poor results in overwintering pupae; natural fluctuations seem desirable. Ample provision must be made for the emergence of adults, if the pupae are in glass, or other smooth-sided containers. A cheesecloth cover, under the lid, and a strip of cheesecloth leading from the bottom to the lid, are very important, to make certain that the emerging insect can climb up to a position where its wings can hang down as they expand and dry. If such provision is not made, the emerging adult will often be ruined. The cheesecloth should be provided soon after pupation, as one cannot
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always tell whether the pupae will go into diapause, or begin to emerge two or three weeks later.
Mr. Christopher Henne of Pearblossom, California, has developed a splendid pupa-container which appears to solve all the problems encountered in keeping pupae alive for long periods of time. Mold and drying out are both prevented, and the pupae remain in excellent condition for months on end. This container includes ample provision for the emergence of adults, and it also has a system for keeping track of numerous different pupae, without getting them mixed up. Mr. Henne has stated that he intends to publish information on this device in the future.
Many very helpful and specific details on caring for larvae and pupae in captivity, are given by Newman (1953).
Preserving My technique follows Peterson (1959), with a few modifications. The technique gives excellent results with nearly all lepidopterous larvae. Many of the colors are perfectly retained, although blues and greens, which are usually due to the color of the body fluid, are always lost entirely; other colors may be altered somewhat. For this reason, it is desirable to keep a notebook of color descriptions to correspond to all preserved specimens. These descriptions should be made from the living larva before it is preserved. It is also worthwhile to include notes on any distinctive habits or behavior (i.e., resting positions; mode of loca-motion; reactions to disturbance; whether or not a nest is built; time of feeding; diurnal or nocturnal, etc.). In describing eggs, color changes should be noted, from the time of oviposition until hatching. In describing a pupa, it should be noted whether or not a glaucous bloom is present, and whether the pupa is capable of abdominal movement, how vigorous this movement can be, etc. If cocoons or earth-cells are constructed, these should be described as to where built, texture, color, and thickness of silk, etc. Many cocoons can be pinned in the dry collection, and are definitely worth saving, as they are often quite distinctive.
The basic solution used in preservation (K.A.A.D.) is as follows: Kerosene — 1 part
( Use ordinary kerosene obtained at service stations—not highly purified kerosene). Glacial Acetic Acid — 1 (or 2) part(s) 95% Ethyl Alcohol - 9 parts Dioxane — 1 part
(Dioxane may be replaced by Iso-butyl alcohol, but more than one part is needed).
Peterson (1959) describes the part played by each ingredient of the solution; knowledge of this makes it possible to modify the basic solution in various ways, in order to achieve good results with all larvae.
I have had such excellent results with the following modified solution, which I call K.A.A.D.I., that I use it almost entirely (for lepidopterous
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208 McFarland: Rearing and preserving larvae Vol.18: noA
eggs, larvae, and pupae):
K.A.A.D. (as given above), using 1 part acid — 12 parts. Iso-butyl alcohol — 3 parts
(Enough must be added to "clear' the' solution, and make the kerosene miscible). Kerosene — 2 parts Glacial Acetic Acid — 1 part
A modification in technique, which gives the best results, is to INJECT all pupae, and any larva over one-half inch in length, using a hypodermic needle, on a two cc. hypodermic syringe. (It is well to have all of these sizes of needles on hand: #27, 26, 24, and 22). It is important that the larva be in the proper condition, before it is preserved. Whatever the instar, it should be nearly "filled out" in that particular instar. Poor specimens result from, those that have only recently moulted a day or two before, or from larvae that are nearly ready to moult.
The larva should first be killed in the solution; a few minutes later, it should be injected (with the same solution) through the anus, to the extent that all the prolegs pop out. If this causes over-inflation, one small puncture with a No. 000 insect pin, in the thin membrane behind the head, will remedy the situation without letting out too much of the injected fluid. Never inject a larva in more than one place, or puncture it with the needle. The injected larva is then returned to the solution, where it should remain for one or more days. If it is a very large larva (size of a tomato sphinx), it should be left in K.A.A.D.I. for about one week. The timing is not of great importance so long as the larva is not taken out too soon. Only experience will show what timing to use; it may vary from 30 minutes, for some eggs and very small or "thin-skinned" larvae, to one week. Little or no damage results from spending more time than required in the solution.
The same solution of K.A.A.D.I. may be used many times, for quite a few larvae, until it becomes a deep yellow-green; most of it should then be thrown out, and new K.A.A.D.I. is added to the preservation-jar, which must have a Bakelite plastic lid that is not subject to reaction with chemicals in the solution.
After sufficient time in the preservation-jar, the larva is removed to 95% ethyl alcohol, in which it is permanently stored. Nothing less than 95% ethyl alcohol should be used, as larvae fixed in K.A.A.D.I. or K.A.A.D. tend to collapse, and they may eventually discolor internally, if the alcohol is weaker than 95%. It is convenient to have "clean-up jars" of 95% alcohol, in which the larvae are first placed; the alcohol in these jars will become green with larval fluids. The larvae are left in these jars for a week or more; finally, they are placed in clean alcohol (95%) for permanent storage, in homeopathic (patent lip) vials. If the larvae have stayed in the "clean-up jar" long enough, the alcohol in the vial will remain clear. Homeopathic vials are far superior to shell vials, for several reasons:
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(1) the stopper fits better; (2) a considerably smaller stopper may be used, reducing surface of stopper exposed to alcohol; (3) an oversize stopper may be forced in (using a No. 1 insect pin to let out air), and this gives a tight seal; (4) the vials don't break easily. Homeopathic vials may be purchased, at a reasonable price, from companies such as Scientific Supplies Company, Division of Van Waters and Rogers, Inc. Ordinary corks are worthless as permanent stoppers, since they eventually break down and shower particles in amongst the specimens, and the alcohol slowly evaporates. Black rubber stoppers are quite unsatisfactory, as they will eventually color the alcohol dark brown, and stain pale larvae; also, these stoppers tend to become stiff, and they may develop cracks. Neoprene stoppers are much more satisfactory. One company which manufactures these gray stoppers is Western Rubber Co., Goshen, Indiana. Neoprene stoppers are low in price and do not discolor the fluid.1 They are very pliable, and give a good, tight seal. The end exposed to alcohol will swell very slightly, but this is of little consequence.
Labels used in vials with preserved larvae are of 100% rag, typing bond paper. This is thin enough not to damage first instar larvae in the vial, yet it is tough and takes black Pelikan (or India) ink very well.
All stages of one life history (eggs - pupae) can be stored in the same vial, when larval size permits. Otherwise, eggs and early instars are placed in one vial, and the larger larvae (with pupae) in other vials. The stoppers in the vials, as well as the locality and determination labels inside, receive the same number that corresponds to the color description, and also to any reared adults in the dry collection. As to size of homeopathic vial, all of the following sizes are useful: 1, 2, 4, 6, and 8 drams. An 8 dram vial will often hold all the stages of a single species, with enough alcohol for permanent preservation. Placing too many larvae in one vial should be avoided; the alcohol will gradually dilute, and then the larvae will begin to discolor (darken) internally. About one year after placing specimens in permanent storage, it is well to go through all the vials once, and replace all green-tinged alcohol with clean 95% ethyl alcohol.
The techniques outlined above will give excellent results with most larvae and pupae, but various modifications must sometimes be employed for special cases. (Experience will show this). Some larvae, especially skippers for example, should be starved for a day or two prior to preservation, and even then they must be thoroughly injected to prevent internal discoloring.
A method for preserving greens and blues is needed, and would be
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iThey are sold only in boxes of five lbs. of one size, and a minimum order is $10.00.
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a significant discovery. For discussion of some other modifications of the K.A.A.D. technique, see Atkins (1958).
References
Atkins, E. L., Jr. 1958. Killing and fixing lepidopterous larvae with a modified K.A.A.D. mixture. Jour. Econ. Ent., 51:740.
Blum, M. S. and J. P. Woodring. 1963. Preservation of insect larvae by vacuum
dehydration. Jour. Kansas Ent. Soc., 36(2) :96-101. Crumb, S. E. 1956. The Larvae of the Phalaenidae. Washington, D. C, (U. S. Dept.
of Agriculture Technical Bull. no. 1135) 356 pp. Forbes, W. T. M. 1930-1960. The Lepidoptera of New York and neighboring states.
Ithaca, New York, Cornell University, 4 vols. Harris, R. J. C. (ed.) 1954. Biological applications of freezing and drying. New York,
Academic. 415 pp. Jones, J. R. J. L. 1951. An Annotated Check-list of the Macrolepidoptera of British
Columbia (Ent. Soc. B. C., occ. paper #1.). Meryman, H. T. 1960. The preparation of biological museum specimens by freeze-
drying. Curator, 3(1):5-19. Newman, L. H. 1953. The Butterfly Farmer. London, Phoenix House. 208 p. Peterson, Alvah 1943. Some new killing fluids for larvae of insects. Jour. Econ. Ent.,
36(1):115. Peterson, Alvah 1953. A manual of entomological techniques. 7th ed. Ann Arbor,
Michigan, Edwards Bros. 367 pp. Peterson, Alvah 1959. Larvae of insects. Part I. 4th ed. Ann Arbor, Michigan, Edwards Bros. 315 p. Rohlf, J. F. 1957. A new technique in the preserving of soft-bodied insects and
spiders. Turtox News, 35 (10): 226-229. Woodring, J. P. and M. S. Blum. 1963. Freeze-drying of spiders and immature insects.
Ann. Ent. Soc. Amer., 56(2): 138-141.
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BOOK NOTICE
FOREST LEPIDOPTERA OF CANADA RECORDED BY THE FOREST INSECT SURVEY. Volume 3, Lasiocampidae, Thyatiridae, Drepanidae, Geometridae. Compiled by R. M. Prentice. Publication 1013, Forest Ent. & Pathol. Branch, Canada Dept. Forestry, Ottawa. 260 pp. (numbered 283-543) including 173 maps (figs. 164-337). 1963. Paper.
This is the third in a series of compilations of the forest Lepidoptera of Canada based on data gathered by the Forest Insect Survey. The general operations of the survey and methods of compiling records were outlined in the first of the series. Volume 1, published in 1958 (Canad. Dept. Agric. Publ. 1034) also included records on Papilionidea, Hesperiioidea, Sphingoidea, Saturniioidea, Nolidae, and Arctiidae. Volume 2 (1962, Dept. Forestry Bull. 128) treated the remainder of the Noctuoidea.
The format of the present volume is identical to that of the previous ones, including records on the distribution, hosts, feeding type, relative abundance, and seasonal occurrence of each species sampled. Numbering of pages, figures, and species is consecutive through the series. Some 260 species of the above listed four moth families, all but 13 Geometridae, are treated in volume 3, bringing the total for the three works to 614. Forest Lepidoptera of Canada is undoubtedly the greatest wealth of information on the biology and distribution of Nearctic Lepidoptera ever brought together in one publication. Indices to insects and hosts are given for each volume. — editor
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