THE EFFECT OF SYNTHETIC PYRETHROID INSECTICIDES ON HONEY BEES IN INDIANA: LABORATORY STUDIES AND A SURVEY OF BEEKEEPERS AND PESTICIDE APPLICATORS

INTRODUCTION

Insecticides are applied annually to a high percentage of the cultivated acres in Indiana. Some of these insecticides are applied in a manner and at a time as to expose non-target organisms, including the honey bee, to direct sprays or residues (Atkins 1979, E.H. Erickson 1983a). Concern over the impact of pesticides on the beekeeping industry has been expressed by leaders of that industry (Ambrose 1983; Atkins 1980; Crane 1983; E.H. Erickson 1983a, B.J. Erickson 1984a, 1984b; Knol 1983; Stevenson 1978, and others). The public is also becoming more concerned about the impact of pesticides on the environment as evidenced by increased regulation pesticide applicators face arid the removal of products from the marketplace (Adkinsson 1971, Pimentel 1980). As many older, more persistent, chlorinated hydrocarbons were removed from use, farmers turned to newer shorter lived insecticides and often found more applications were necessary to achieve acceptable control.

One class of insecticides that contains many of the newly registered insecticides is the synthetic pyrethroids. As a group, the synthetic pyrethroids are loosely related by their chemistry and mode of action on the target pests. Within the group are a wide range of products that have diverse target pests. Most of these products are characterized by their relatively low mammalian toxicity and their effectiveness against invertebrate pests at low doses (Sine 1988). In Indiana, these products are being utilized in the pest management programs of a growing number of corn, soybean and alfalfa farmers.

Some synthetic pyrethroids are reported to be quite safe to honeybees in some areas of the United States (Atkins 1979, Johansen 1983, Moffet 1982, Stoner 1984). Early evidence indicates that the toxicity of some synthetic pyrethroids to honeybees may be greater in Midwestern areas than in warmer more arid parts of the country (B.J. Erickson 1983; E.H. Erickson 1983b,1984; Smart 1982). The fact that the toxicity of some synthetic pyrethroids is inversely related to temperature (Georghiou 1964, Morton 1979) and may be important in a contaminated honey bee colony’s ability to overwinter in the midwest. It was also evident from early studies that the toxicity of this group to any particular species was very diverse (Atkins 1981, Moffet 1982, Smart 1982). For example, some of these products are highly toxic to mites and are used as miticides (Herbert 1988, Witherell 1988), while others are so safe to mites as to actually increase their population (Flaherty 1981, Flint 1985).

As farmers have become more specialized producers of a declining range of crops, the direct importance of bees as pollinators on the farm has also been declining. While some studies indicated that soybean yields might benefit from insect pollination (Abrams 1978, Erickson 1978, Mason 1979), other crops such as corn, alfalfa grown for hay and wheat require no insect pollination. As these crops captured an increasing proportion of the acreage, the number of nectar producing plants plummeted. The decrease in forested acres, the intensive planting of non-nectar producing plants such as fescue and crown vetch on roadsides and in pastures, the intensive use of herbicides in cropland and the increasing urban demand for land, seriously reduced the nectar resources available to bees. This combined with the low world honey price over the past several years, has driven nearly all commercial beekeepers from many parts of Indiana.

Most of the remaining beekeepers in Indiana are hobbyist or part-time beekeepers who keep bees for pleasure as well as profit. For these beekeepers, the impact of pesticides on their bees is a highly-charged, emotional issue. Much misunderstanding exists between beekeepers and applicators and there is considerable misinformation on both sides. Since discontinuation of the federal governments Beekeeper Indemnification Program, there has been very little effort to evaluate, document, or record reports of pesticide poisonings of honey bee colonies, unless litigation was instituted or threatened (Coleman 1979, Pimentel 1981). It is generally believed, but undocumented, that beekeepers have overestimated the severity of the problem, while pesticide applicators have underestimated the extent of the problem.

In Indiana, application of those products which have been classed by the United States Environmental Protection Agency (EPA) as “Restricted Use Pesticides is regulated by the State Chemist’s Office. This office has the responsibility for enforcing the laws and regulations relating to pesticides including pesticide applicator testing and certification.

Because many individuals who apply pesticides may never use a restricted product, many private applicators are not required to be trained or show competency in pesticide use. The Indiana Cooperative Extension Service and the Office of the State Chemist have worked together to train pesticides applicators in the safe handling and application of pesticides, as well as safety to the environment. Pesticide applicators are further divided into those who apply pesticides for hire and those who use the products only in conjunction with their own crop production operation. These groups are referred to as Public Pesticide Applicators and Private Pesticide Applicators, respectively.

Among the information which is required for certification is knowledge of the safe handling and use of pesticides, including their toxicity to non-target organisms such as the honey bee. The issue of honey bee poisoning is a complicated one and can not be covered in depth during applicator training due to time constraints.

The factors which determine the extent to which a given colony of honey bees will be affected by the application of a pesticide to a given field are complex (Atkins 1981, Johansen 1977, Lieberman 1964, Quatitlebaum 1983). The single most important factor is the number of bees from a particular hive that are foraging in the treated area (Nowakawski 1982). This is influenced by many factors such as attractiveness of the crop treated, presence or absence of blooming weeds in the area, distance from the treated field to the colony, strength of the colony, weather, needs of the particular colony, genetic make-up of the colony, etc (Atkins 1977, B.J. Erickson 1983a, Mayer 1983, Mayland 1970, Smirle 1987, Ross 1981, Wailer 1984).

In addition to factors relating to the honey bee colony, factors relating to the pesticide such as the active ingredient, the formulation, the time of application, the method of application, the weather conditions during and following application, etc., will all influence the extent to which a given colony will be affected. There is the additional complicating factor that some pesticide products may cause no observable damage at the time of application, but may cause delayed mortality of the overwintering colony during a period of greater stress.

The synthetic pyrethroid insecticides are of particular concern in this regard due to the inverse relationship between their toxicity and temperature (Yu 1984). Lehner (in press) has shown that the toxicity of permethrin to bees dramatically increases at 20 degrees C over the toxicity at 26 degrees C. Delayed mortality may often not be detected or identified as a result of earlier pesticide exposure. The insecticide stored in the hive may not singularly cause colony mortality, but may act in conjunction with other factors to increase the stress on the hive and cause a decline of the population. This decline may or may not be reversed by the colony as weather and other conditions improve, depending on their reserve strength and size of the initial population.

An understanding by both beekeepers and pesticide applicators of the factors that influence poisoning of colonies of honey bees by pesticides is critical to establishing a situation in which the two groups can operate without conflict. Because this is such an emotional issue, it is often difficult to separate emotion from fact when discussing this subject with either side. Given the complexity of the problem and the limited resources available to try to deal with the situation, a multi-faceted approach to the problem was undertaken. This included laboratory work to examine some the the most critical questions relating to synthetic pyrethroids and bees as well as an examination of the groups of people directly involved; that is beekeepers, farmers and public pesticide applicators in Indiana.

THE EFFECT OF SYNTHETIC PYRETHROID INSECTICIDES ON HONEY BEES IN INDIANA: LABORATORY STUDIES AND A SURVEY OF BEEKEEPERS AND PESTICIDE APPLICATORS

ABSTRACT

CHANEY, WILLIAM EUGENE. PhD., Purdue University, August 1988.
The Effect of Synthetic Pyrethroid Insecticides on Honey Bees in Indiana: Laboratory Studies and a Survey of Beekeepers and Pesticide Applicators.

Major Professor: C. Richard Edwards.

Insecticides are an important component of the row crop production system in Indiana. Concern for the safe use of these products has lead to a system of regulating the application of pesticides that is designed to protect the public, the environment and the applicator. One non-target organism that is affected by some pesticide applications is the honey bee. Because of its social nature, the impact of pesticides on bees is sometimes expressed as detrimental effects on the colony to which the exposed bee delivers her contaminated nectar or honey.

This study looked at several aspects of the honey bee/pesticide problem, including one class of insecticides about which there is controversy concerning their impact on bees. This class is the synthetic pyrethroids. These studies found that the relative toxicity to adult bees of the four products examined was: permethrin> flucythrinate > fenvalerate > fluvalinate, in decreasing toxicity. The toxicity of these products was also shown to increase at 18 degrees C and 12 degrees C as compared to their toxicity at 25 degrees C. These are temperatures in a range which might be experienced by bees in a colony in Indiana during the winter.

This study also demonstrated that no synergism or antagonism was seen when permethrin and fluvalinate were fed to adult bees together with carbaryl, paraquat or mancozeb. This study did demonstrate that some colonies were more resistant to permethrin and carbaryl than others and that this resistance was related to the race of the queen heading the colony.

Beekeepers, public pesticide applicators and private pesticide applicators were surveyed to examine their knowledge of and attitudes toward the poisoning of honey bee colonies by pesticides. The response rate was not significantly different among the groups. The mean response rate was 75%. Less than 10% of the beekeepers and none of the applicators reported any knowledge of specific incidents in which bees were poisoned by pesticides in 1986. Both the beekeepers and the applicators were concerned about this issue and both groups indicated a willingness to take specific actions to attempt to prevent future poisonings. Each of the three groups showed a poor level of knowledge about pesticides as they relate to bees and about integrated pest management.

A Thesis Submitted to the Faculty of Purdue University by

William Eugene Chaney

In Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy.

August 1988

ACKNOWLEDGMENTS

I wish to express my appreciation to my major professor, and friend, Dr. Rich Edwards, for his support in my graduate work, my professional career, and my personal life. I would also like to thank Dr. Eldon Ortman for his support as head of the Department of Entomology. A special thanks to Dr. Marlin Bergman for his encouragement and to Dr. Harry Potter for his help in the survey portion of this work. My gratitude also to Dr. Virgil Anderson who made a statistical difference in my graduate education and to Dr. Tom Jordan for stepping in at the last minute.

Most importantly, I would like to thank several outstanding individuals without whom this work could have never been done. They are, in no particular order: Carl Geiger, Terry McCain, Suzie Appel, Mark Johnson, Mike Gale, Steve Mroczkiewicz and Connie Holderfield.

ABSTRACT
INTRODUCTION
CHAPTER ONE - RELATIVE TOXICITIES AND TEMPERATURE
CHAPTER TWO - PESTICIDE INTERACTIONS
CHAPTER THREE - INTERHIVE VARIABILITY

Bee Pathology - pages 171-173

THE TRUTH ABOUT VARROA IN BRAZIL
GONCALVES, L. S.; DE JONG, D. (Brazil)
MORSE, R. A. (U.S.A.)

Varroa is currently recognized to be the greatest problem for apiculture worldwide. First described in Asiatic bees Apis cerana, in Indonesia, by the Dutch researcher Oudemans in 1904, the mite turned into a problem for Brazilian bees, Apis mellifera, in the 1970s. Transfer of the pest from A. cerana, whose infestation is of little economical importance, to A. mellifera occurred when beekeepers carried honeybees to Asia. Today Varroa is found throughout Asia, in most of Europe, in the Northern region of Africa, in several countries in the Middle East and in the following South American countries: Paraguay, Brazil, Argentina, Uruguay Peru and Bolivia.

In 1971, infested hives were brought to Paraguay from Japan. Soon after, in 1972, an apiculturist from Sao Paulo took some of these Japanese bees to the region of Jundiai, thus initiating infestation in Brazil. However, infestation was first detected in 1978 when the pest had spread considerably and there was no way of eliminating this “new” enemy of Brazilian apiculture. At present, Varroa can be found in 18 Brazilian States, from Rio Grande do Sul to Piaui, so that it is too late to avoid transporting hives from a State to another.

Can we blame the Japanese, or other peoples, or even our own beekeepers for spreading so much these bees without taking precautions against possible consequences such as the introduction of Varroa? In a way, we can. All those who work in apiculture should he aware of the possible consequences of their carelessness, especially those involved in ”migrant apiculture” at the international level. A period of quarantine is always recommended to avoid the entry of pests into the country. In 1971, Varroa was little known and the problems of hive mortality, which were already occurring in Russia, were not properly communicated to the Western world. In Eastern Europe also, Varroa was present 4 to 6 years before being detected. Not even West Germany, where apiculture is well organized and controlled, succeeded in containing Varroa.

Research on Varroa in Brazil

The Department of Genetics of the University of Sao Paulo, Campus of Ribeirao Preto, started a research program on Varroa in 1979 under our direction and with the financial support of the Foundation for the Advancement of Science in the State of Sao Paulo (FAPESP) and later of the National Research Council (CNPq). ln 1980, a collaborative program between USP and the Department of Entomology of Cornell University was started, with the financial support of the National Science Foundation (NSF) and later of the Department of Agriculture, USA (USDA). As part of this program, Dr. De Jong came to Brazil in 1980, where he is currently involved in research.

It was soon observed that Varroa had spread considerably but with low indices of infestation. The mite showed no symptoms to the beekeepers who did not know it, since at the beginning of infestation Varroa is present in very reduced numbers in hives, although it rapidly transfers from hive to hive through new field bees which get lost during the first recognizance flights and through drones, which tend to enter other hives.

What is Varroa?

Varroa is an acarid which attacks honeybees. It is a relatively large brown mite 1-mm long and 1.6-mm wide. Varroa feeds on bee “blood” (hemolymph). Normally, only adult females can be seen. The other female stages (nymphs) and males are found only in bee brood cells and are smaller and white. These also feed on bee “blood” (except for adult males which do not feed on anything). When compared with other types of pests, Varroa, reproduces slowly. Under ideal conditions, the mite leaves on average one to two new female descendants per cycle in worker bee brood. Without taking into account mortality or migration to other hives, an infestation starting with a single female may take approximately 400 days to include 5000 acarids. At the same time, many of these females may be moving to other hives, and therefore it would take even longer for an initial infestation to reach a damaging level. Obviously, if infestation started by introduction of well-infested hives into a beekeeping unit of a migratory type, proliferation becomes more rapid. Also, Varroa multiplies more rapidly at the time of drone rearing, with a single female capable of producing 2 to 4 new females in a drone cell.

Our studies have shown that Varroa reduces the weight and mean life of bees that were infested during development. When infestation is low, however, the beekeeper does not detect this damage. When a single larva is attacked by 5 or more acarids it can still survive, but its body, and the wings in particular, will be visibly damaged. However, very few bees are attacked by so many acarids, and when this happens they are thrown out by the other bees, so that the beekeeper will not see the problem.

Hive mortality caused by Varroa

Millions of hives have already died in Europe because of Varroa. Tens of thousands have died in Argentina. In many regions of the world it is impossible to keep bees without treating them. In Brazil thus far, no hive death caused by Varroa has been reported. even though the mite reached Brazil before being introduced into Argentina. A completely unexpected phenomenon has occurred, since greater infestation was expected in our climate, which permits Varroa to reproduce throughout the year, in contrast to countries with cold climates, where Varroa does not reproduce in winter.

Fortunately until now we escaped relatively well, since our levels of infestation with the pest are low, whereas they are very high in the rest of the world. However, Varroa still causes serious concern. Even though the level of infestation is low and does not cause death of hives, there is still damage, because all hives in the country are progressively becoming infested. Infestation causes little damage per hive, but if we sum all the hives, great waste occurs in our apiculture.

We can estimate the damage caused by Varroa as follows: our studies have shown that the mean life of bees infested with Varroa during development is reduced by half. Thus, for each two bees affected, we actually lose one. If 2% of worker brood is infested, the population of the hive will be reduced by 1%. We may estimate that his would cause a 1% reduction in honey production. Since national production in 1984 was estimated at about 30,000 tons, the reduction would be approximately 300 tons which, at the cost price of Cr$9,500.00 per kilogram, would cause a loss of about 3 billion cruzeiros (about 300,000 US$). The ideal would be for all beekeepers to be able to treat their bees to save them and to guarantee honey productioin. However, the solution is not so simple, since so far there is no adequate pesticide for the conditions prevailing in Brazil. Either the treatment is too expensive, or is harmful for the bees, or it contaminates the honey. If one opted for treatment with chemical products, such treatment should be applied every year, at a cost that would be higher than the loss caused by Varroa.

It has been well known since Rachel Carson first wrote “Silent Spring” in the early 1960s that agriculture has been getting deeper into trouble with continued pesticide use.

Today within our beekeeping industry, this has never been more apparent. Yet, strangely, many beekeepers are uneducated in the field about what pesticide resistance is and how it escalates into a pesticide highway to hell and eventual colony destruction. In the past in the 1960s and 1970s forward, it was often described as a pesticide treadmill, because once you are on it with your agricultural field management, it is virtually impossible to wean your self off. Herein therefore is the danger, namely a growing resistance, creating a growing chemical dependency, requiring stronger and stronger treatments of various dopes.

Someone sent me a perfect example, when they wrote, “I have the gut feeling this is how it is as when you have treated with acid, (referring to chemical resistance we had corresponded about with honeybees and parasitic mites), you get a feeling the colony is like a magnet on mites or starts producing more. I have an example from a beekeeper, he treated and treated different kinds, different acids, culling dronecomb, Apistan, everything, and saved all mites and counted them. Though he treated like a maniac, he managed to produce 10,000 mites from that colony in a year. And it was in a miserable state the next spring, but still alive, which surprised me. After all these chemicals. Amazing.”

Well, this could be described as a perfect example of a chemical treadmill in action going full steam with a high degree of resistance to chemicals by the parasitic mites, now totally - out of control!

For understanding, Pesticide resistance or “pest” resistance should be simply stated, - that some insects within each given species are naturally more resistant to certain chemicals. You never can kill 100% of the little trouble makers with any given dope treatment. There is always an exception. Has to be or evolution would stop!

Within our beekeeping community as those treating with various dopes try to kill parasitic mites infesting their colonies, most are not aware that as the dopes do their job and make the susceptible parasitic mites die, the survivors multiply, passing their resistance onto the next generation for that particular level of doping (strength of chemical used).

When the pesticides are perceived to “no longer control” (beekeeper sees a growing body of prolific mites) at normally recommended rates (strength of chemical used), a pest resurgence occurs, when the parasitic mites killed by the dopes return in larger numbers.

Basically, what this is, is the reproduction of those little trouble makers, the exceptions, the dopes could not kill in the first place, now reproducing without chemical effect, their next generations. We say the pesticides no longer control at normally recommended rates, instead of saying we need more killing power to now go back and refight the exceptions that were stronger in the beginning and really needed more dope to finish the job, but now the insect, somehow knows what to expect; and it also requires a higher dosage. Our problem is figuring out how much and of course, here we go again trying to get another 100% kill, which we know is technically impossible. Consequently a circle of treatment or a pesticide treadmill is created.

Now during this treatment, some of the parasitic mites will develop what is called cross-resistance. This is basically where resistance to one chemical means resistance to a second chemical with a similar mode of action (method of killing) as the first. Multiple resistance also is known to occur, where there is resistance to several classes (different chemical groups). This is now currently accomplished, by beekeepers all using different types of treatments within a given area and then as the honeybees co-mingle in the field or drift from colony to colony, the mites transfer rides on the backs of the bees themselves and cross mate, passing each others resistance on to the others linage.

Now to add insult, beekeepers need to understand that depending upon the type of resistance (type of dope used) and the species of parasitic mite (trachael or varroa or ?), resistance tends to last in the absence of the dopes when control measures are stopped. What this means, is that the breeding accomplished through survival (surviving the various dopes thrown upon their bodies) is now considered inherent within the linage of the parasitic mites.

Now, as beekeepers use stronger and stronger dopes other problems begin to set in to complicate the already growing bad situation created. Many times other problems created involve not only pest (mite) resurgence, but also the creation of secondary pests/insects now becoming serious primary pests not known to effect colonies i.e. beetles, ants, earwigs, moths, and also secondary diseases.

This happens because when we attempt to kill what we consider our primary pests, namely parasitic mites, we also inadvertently also kill their natural enemies that would help to keep them in balance around our hives. Two things normally happen here. Either the natural enemies of our pests are killed or they leave the area since their food source is no longer available. This leaves opportunity for the treated surviving parasitic mites to reproduce before their natural enemies return (other insects, or birds, etc.).

Secondary pests become serious primary pests when their natural predators are killed. The whole class of mites is very widely studied for this vary reason for the havoc caused when dopes kill their natural predators. This adds to the pesticide treadmill. Then the corresponding treatment of the secondary pests i.e. beetles, moths, ants, etc., add to the problem of our mites and our ability to contain their damage. Look at beekeepers here in the USA currently treating for parasitic mites, now also being forced to treat for beetles with chemicals, now used not only within the colonies, but now all around the colonies on the ground. This is nothing more than a serious speeding up of the treadmill to disaster because you have increased the scope and width of the playing field for chemical management.

Add to this now, besides external dopes, internal dopes like oxytetracycline that blow out the bees internal gut for beneficial digestive bacteria and you now create not only eating disorders, but internal susceptibility to various secondary diseases and shorten the life of our poor honeybees at the same time.

Yet to stop the dopes both external and internal is pure hell. For now there is no natural backup help to be found, the bees are in a weakened state not able to digest natural food, they, for the most part can no longer defend themselves, and then we wonder why colonies collapse.

What are we left with? I would say a lot of empty equipment growing in quantity each and every year as the treadmill, now worldwide accelerates, persistent residues in soil and living tissue for those stupid enough to eat this wholesome food we still call honey, and more and more decreased pollenation service sure to effect our global food supply in the future. Yet, if I may be so bold to state - very few are willing to do what it takes to go back to full biological to correct the problem. But in the end I really feel in my heart they may be forced to, and what a price we are all going to pay as a worldwide global community.

- Dee Lusby

Science Bulletin 220
UNION OF SOUTH AFRICA
DEPARTMENT OF AGRICULTURE AND FORESTRY
(Entomological Series 3)
1940

A. E. LUNDIE, Ph.D.,
Research Apiculturist Division of Entomology, Pretoria

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INTRODUCTION

It is surprising that the insect to be described in this paper, although it is so common and can be so troublesome to the beekeeper, has not been mentioned previously in our apicultural literature. The absence of any mention of this trouble by beekeepers contributing to our journals is but another reminder that South African apiculture is still in its very infancy.

It is suggested that this insect, Aethina tumida, should be called “the small hive beetle” to distinguish it from another and much larger beetle, Hyplostoma fuligincus, which is also to be frequently found in bee hives in South Africa. Although the mature beetles are black, it may be well to avoid this adjective in any descriptive title to be given to the insect, because some beetles, on emerging from the ground, are a very light brown, and the period during which they turn from light brown to a dark brown and finally black is variable. However, the majority of these beetles after reaching maturity do not become active till a fairly high degree of pigmentation has set in.

The writer first became acquainted with this small beetle in 1924, shortly after hiving his first swarm of bees in Pretoria. A nucleus from this swarm was set aside for increase, but a later visit showed that the bees had absconded, and that the combs had become a seething mass of “worms”, which were easily recognized as beetle larvae.

Various facts about this insect and its habits were gleaned from time to time in practical work in the apiary, but it was not till 1931 that the writer, due to curtailment of his itinerant work, had the opportunity of commencing a detailed study of the insect. After further interruption the study was resumed in January 1938 and this paper gives the results of the investigations to date.

Our early entomologists received a few queries from beekeepers on an insect which was undoubtedly Aethina tumida, but in the absence of any detailed work on this insect it was suggested then that these beetles were probably not a pest, and that they were merely associated with the bees for the shelter and warmth which the hive and cluster afforded.

The first South African record of the insect is by Mr. R. H. T. P. Harris, who submitted specimens for identification from Durban in 1920.

Aethina tumida was first named and described by Andrew Murray in 1867 in the “Annals and Magazine of Natural History”, London, from two specimens which were sent to him by the Rev. W. C. Thomson from Old Calabar on the West Coast of Africa, but no mention is made of the insect being associated with honey bees in any way. This reference indicates that Aethina tumida must be widely distributed over the African continent. The writer has failed to find any other reference to the insect in the entomological literature available to him.

Aethina tumida belongs to the family Nitidulidae, about which W. S. Blatchley remarks that “The name Nitidula applied by Fabricius to the typical genus, is very inappropriate for the family, since it literally means shining or elegant, whereas the great majority of the species are clothed with a fine pubescence which does not permit of their to any great extent”. This fine pubescence is present in Aethina tumida.

The habits of this family were picturesquely described by Murray in the “Transactions of the Linnean Society”, London, in 1864, as follows:-

“The chief function of this family is that of scavengers. Their main business is to clear off decaying substances from the face of the earth, especially those minute and neglected portions which have escaped the attention of other scavengers whose operations are conducted on a larger scale. We may characterize them in one point of view as retail scavengers. They are, so to speak, users-up of waste materials. After the beast of prey has satisfied his hunger on the animal he has slain, after the hyena and the vulture have gorged themselves on its carrion, after the fly with its army of maggots has consumed the soft parts, after the burying beetles and the Silphidae have borne their part in the clearing away and when nought but the bones remain, then come the Nitidulariae to go over what they have left, to gnaw off every fragment of ligament or tendon and to leave the bones as nearly in the state of phosphate of lime as external treatment can. In another point of view, however, their employment is wholesale and wide enough. They conduct their operations all over the world, their branches extend into the most remote district; the materials with which they have to do, although mere waste, have no other limit to their variety or their number than the organized substances found on the surface of the globe. As in all great establishments, too, the principle of division of labour is carried to a great extent. Each different kind of substance has a different member of the firm told off to take charge of it. One species confines itself to rotten oranges, another to bones, a third to putrid fungi, a fourth to decaying figs. Decaying wood, decaying bark, decaying flowers, decaying leaves, all furnish distinct employment to different species. They are not all scavengers, however. Many pass their lives in flowers; others feed upon fresh victuals; and Mr. Frederick Smith, of the British Museum, has, while I write, brought to my notice a species of Brachypeplus (B. auritus) which he has received from Australia, in a wild bee’s nest, where it feeds, both in the larva and perfect state on the wax and honey.”

In discussing the Trigona or stingless bees of Australia in his book “A Cluster of Bees”, Tarlton Rayment mentions three beetles, Brachypeplus planus Er., Brachypeplus meyricki Blkb. and Tribolium myrmecophilum Lea and their association with these bees, but they are evidently not associated with honeybees in the same way as Aethina tumida, because he reports on the more abundant species B. planus, “Beetles placed on the combs of hive bees were immediately carried off by the workers”. Honeybees cannot eject A. tumida beetles so easily. These beetles can invade strong colonies of honeybees as well as weak ones with equal impunity.

These two references, both from Australia, are the only ones the writer has been able to find, which report an association of beetles and social bees which bears some similarity to the association of A. tumida with the honeybee in Africa.

Although A. tumida is not a major pest, there are localities where, and seasons when, it assumes such importance that it becomes almost as serious as the compolitan wax moth, and its control requires an equal vigilance from the beekeeper.

The absence of any reference to this beetle in our apicultural literature is probably due to the fact that most beekeepers who have been troubled by the larvae of this beetle have mistaken these larvae for wax moth larvae; and to add to their confusion combs are often infested simultaneously by the larvae of these two insects. The morphological differences between lepidopterons and coleopterous larvae are not generally known by laymen.

Although the beetles may be found anywhere in the hive, their favourite rendeavous seems to be the rear portion of the bottom board, where they probably escape to some degree the attention of both the bees and the beekeeper. Here not only are the beetles out of the maelstrom of traffic to and from the hive, but probably they can also secure their food, with a minimum of interferance from the bees, from the pellets of pollen that fall from the cluster of bees above them. The beetles, however, may be seen, immediately the inner cover is removed from the hive, lurking in the cavity behind the metal rabbets or in cavities in any burr-comb the bees may have built between the top bars of the frames and the cover of the hive.

When the frames of the hive are examined, these beetles may be seen running along the surface of the comb to disappear for a moment in a cell before emerging again to continue their scramble for a “safer” hiding place, perhaps in another cell nearby, where they may remain motionless and so escape the attention of the observer.One may also see a bee “close with” one of these beetles, curling its abdomen around the beetle in a vain endeavour to penetrate the hard chitinous armour of he intruding beetle, or the bee may be fortunate in getting a good hold of the beetle in this struggle and, taking flight, it “jettisons” the invading beetle at some distance from the hive. However, this does not appear to happen very frequently, for many beetles can live for long periods of time, even in strong colonies, with relative impunity.

THE NATURE OF THE PEST.

The small hive beetle is a scavenger, which may be likened to the cosmopolitan wax moth in many ways, but fortunately it is not nearly so destructive to the combs.

Just as the wax moths begin their ravages in combs in storage, or in weak colonies, so does the small hive beetle become a nuisance to the beekeeper. Any factor which so reduces the ratio of the population of a colony of bees to its comb surface that the bees are no longer able to protect this comb surface adequately is a precursor to the ravages of both the wax moths and Aethina tumida. Such factors as incorrect supering of the hive, excessive swarming, long standing European foulbrood, pilfering of some of the honey of the hive by thieves, who may pour water over the bees or use excessive smoke in obtaining their spoils, may result in a heavy infestation of Aethina tumida larvae, before the beekeeper is able to improve the condition of the colony.

The following are some of the principal occasions when a beekeeper may experience some trouble from this insect. Combs of honey that have to stand in the honey-house for a period before or after the extraction of the honey, are liable to become “wormy”, especially those combs that contain a certain amount of pollen. Cappings which are invariably set aside at extracting time by the beekeeper to be worked at a later date are liable to become “wormy” before they are melted down into cakes of wax. Honey left over Porter bee-escapes for several days before its removal may develop the larvae repidly as a result of the additional warmth which this honey gets from the colony of bees below it.

The larvae will pierce the cappings, side walls, and mid-rib of newly made or other relatively delicate comb, causing the honey to “weep” badly (Fig. 1 and Fig. 2), but old combs with several generations of cocoons can withstand heavy infestations well (Fig. 4) and can be used again in the hives after the gummy mixture of honey and larval excrement has been washed off with water under some pressure, such as that from a garden hose.

There are two characteristic conditions of the larval infestation which depend upon the relative abundance of honey and pollen in the infested area of the comb. When this area contains a small amount of honey and the larvae are feeding on the pollen mainly, their faeces have a dry “shredded” appearance, and the larvae themselves are a bright, dry, clean white; but when the honey being worked by the larvae is more abundant, this at first becomes discoloured, due to the faeces which the larvae void in the honey; then it becomes thin and ferments with a very characteristic odour, somewhat like that from decaying oranges. This odour in a honey room is usually the first warning to a beekeeper of the presence of active larvae in his supers. As the fermentation progresses, frothy bubbles ooze out of the cells of the comb (Fig. 1) and the “honey” falls to the bottom board where, in the case of an old infestation, it may accumulate sufficiently to run out of the entrance of the hive to the ground, or collect to form a layer an inch or more thick should the entrance become blocked or the hive bottom slope be to the rear of the hive. In this case the larvae become so thoroughly covered by the fermenting honey that they present an unpleasant slimy appearance, and when they begin to migrate away from this mixture, they leave trails of it behind them, discolouring everything over which they crawl.

With further fermentation and drying, the mixture of honey and larval excrement becomes sticky, and still later shrinks to a granular or somewhat spoungy mass, which can neither be scraped nor washed off easily from the bottom board. The full-grown larvae leaving the hive through any crevices large enough to give them egress on their way to the ground to pupate may carry a small proportion of this sticky mixture to the outside of the hive. Tracks of the mixture may be left in such quantities that in a heavy infestation even the outside of the hive may become quite badly discoloured [Fig. 5 and Fig. 6 (a)] by the hosts of migrating larvae on their exodus from the hive.

A perusal of Fig. 6 (b) will give some idea of the number of larvae which can develop in a few honey combs. This illustration represents a few of the dead larvae collected from the concrete floor of a honey room, where an infestation of some of the honey combs had occurred. These larvae died shortly after reaching maturity, having failed to find a suitable place on the hard concrete surface in which they could pupate.

Apiaries that have been established for a number of years are more likely to harbour a larger number of A. tumida beetles than recently established apiaries. Once a colony or a number of colonies in an apiary have retrogressed so far that these beetles have been able to breed in considerable numbers, other and normal colonies in the same apiary will harbour a greater number of these beetles, and there will be the danger that any supers from such an apiary will develop “wormy” combs rapidly, soon after they are left in storage in the honey-house.

One of the riddles of beekeeping is the total absence of American foulbrood in South Africa. This disease is prevalent in the Mediterranean countries and seems to be present in Northern Africa; but just why should Southern Africa be free of this disease, when conditions seem so ideal for relaying it down the length or “backbone” of the African continent? Perhaps in the warm tropics the rapid work of scavengers, of which the wax moths and Aethina tumida must play an important role, accounts for the absence of this foulbrood in Southern Africa.

DISTRIBUTION OF Aethina tumida.

In an attempt to get some information on the distribution of Aethina tumida in South Africa, a questionnaire on this insect, accompanied by specimens of the beetle, was sent to forty-four beekeepers of long standing. Only eleven beekeepers of the thirty-one that responded to this questionnaire showed that they were familiar with the beetle or its larvae. Ten of these beekeepers live in the low-veld or warmer areas of the Transvaal, and in the coastal areas of Natal and the Cape Province. One beekeeper on the Transvaal high-veld, who at first reported that he had never seen the beetle before, sent in specimens from his apiary at a later date. Beekeepers of very long experience in the Western Province and the Cape Midlands were not familiar with the beetle. Perhaps the climate and the nature of the soil militate against a rapid development of the beetle in these areas.

The presence of the beetle in Old Calabar, on the West Coast of Africa, suggests that A. tumida is widely distributed over the African continent, and in the absence of definite records it may be assumed that the beetle will be found in any of the tropical and subtropical regions of Africa.

CONDITIONS UNDER WHICH THE LIFE HISTORY OF Aethina tumida WAS STUDIED.

A knowledge of the inability of A. tumida to live long without regular supplies of fresh water and of the humidity requirements of the soil for the pupal period, gleaned from the 1931-32 study, enabled the writer to make more rapid progress in the 1938 study of this insect.

At first the full-grown larvae obtained from a heavily-infested hive were placed on damp soil in three types of containers:-

1. Small tin boxes 1-3/4 inches in diameter and 1-1/2 inches high with transparent lids.
2. Larger tin boxes, the diameter and height of which were about 3 inches to 3-1/4 inches, with loose-fitting metal lids.
3. Glass battery jars about 4 inches in diameter and 6 inches high, covered with two sheets of transparent paper or muslin held in place by a strong elastic band.

There was a high mortality of the pupae in the small tins, due to the small volume of soil present and the free passage of air through the junction of the transparent material and the metal rims of the lids, which dried the soil rather rapidly. In spite of the larger volume of the glass jars and the use of paper covers to retard evaporation, the soil in these containers also dried out too rapidly.

The larger tin boxes proved to be the most satisfactory and were used throughout the greater period of this study. The soil was sifted through a piece of perforated metal with holes 1/16 inch in diameter and remained moist long enough for several generations, without any addition of water. The degree of moisture which was maintained in these tins may be judged by the ease with which several specimens of the common earthworm (Lumbricus sp.) grew to a length of about three inches in the soil in some of these tins and were kept in this way with no further addition of moisture for several months.

All the tins were kept on a table in an unheated room some 12×12x12 feet in extent and having a window (3×6 feet) on the north side of the building. The room was used as the writer’s office. Its temperature would approximate that of any medium-sized honey-room used by beekeepers in extracting and storing their honey.

The adult beetles were kept in the larger tins. They were removed daily to a clean tin containing fresh food and a clean piece of cotton wool soaked in water, except at the week-ends, when the beetles would be two days with one lot of food and water. The food supplied was a mixture of honey and pollen, thoroughly worked together to form a thick paste. The larvae were also fed on this mixture, but no water was supplied to them. The ease with which A. tumida can be bred in tins or petri dishes and the longevity of the insect, would make it a very suitable one to breed for general laboratory purposes or for museums exhibiting live insects.

Insecticides, Acaricides and Ovicides
BY W. T. THOMSON

COUMAPHOS, ASUNTOL, BAYMIX, CO-RAL, DIOLICE, MELDANE, RESITOX, UMBETHION, NEGASHUNT, PERIZIN 0,0-diethyl-0-(3-chloro-4-methyl-2-oxo-2H-l-benzapyran-7-yl)
phosphorothioate

TYPE: Coumaphos is a systemic, organic phosphate livestock insecticide.

ORIGIN: BayerAG in Germany, 1958. Licensed to be sold in the U.S. by Mobay Animal Health Div.

TOXICITY: LD50-13 mg/kg. May cause eye and skin irritation.

FORMULATIONS: 25% WP, 1 and 5% dusts, 4% pour-on, 3% spray foam. 4.2 EC, 1 1.6% EC.

PHYTOTOXICITY: Generally not applied to plants.

USES: Used on beef cattle, dairy cattle, sheep, dogs, goats, swine and horses.

IMPORTANT PESTS CONTROLLED: Grubs, flies, lice, ticks, keds, poultry mites, screwworms, mosquitoes, and others.

APPLICATION:
1. Backline treatment-Apply .5 oz/100 lb body weight for grub control.
2. Spray-Apply after heel fly season has passed at 250 psi or more pressure. Apply approximately 1 gal of the diluted spray per animal. Wet the skin, not just the hair.
3. Dip-Agitate the tank thoroughly prior to use. Repeat as necessary. Maintain adequate concentration in the vats.
4. Spot treatment-Use in infected wounds for screwworm control. May also be applied as a dust.
5. Backrubbers-Place backrubbers where animals travel daily. Refill as needed.
6. Cattle grubs-Treat at least 6 weeks before the expected appearance of the grubs in the back.

PRECAUTIONS: Do not spray animals in a confined, unventilated area. Do not apply to sick or stressed animals or animals less than 3 months old. Do not dip overheated animals. Do not treat within 10 days of shipping, weaning, vaccination, etc. Do not use before or after the application of natural or synthetic pyrethrins or compounds used to synergize them. Cattle on a fattening ration may be more subject to organic phosphate poisoning than animals on pasture or maintenance feed. Do not mix with other insecticides nor use in conjunction with oral drenches or other internal medicines. Toxic to birds and fish.

ADDITIONAL INFORMATION: 10-20 day protection from screwworms can be obtained. Used to control fly larvae in poultry manure. Systemically controls cattle grubs in cattle. Used to control insects on humans.

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FLUVALINATE, KLARTAN, MAVRTK, SPUR, YARDEX, APISTAN
(RS)-alpha-cyano-3-phenoxybenzyl(R)-2-[2-chloro-4-(trifluoromethyl)anilino] -3-methylbutanoate

TYPE: Fluvalinate is a synthetic-pyrethroid compound used as a selective contact and stomach-poison insecticide.

ORIGIN: Zoecon Crop., 1980. Now being marketed by Sandoz Crop Protection.

TOXICITY: LD50-261 mg/kg. May cause eye and skin irritation.

FORMULATION: 2 lb/gal flowable.

PHYTOTOXICITY: Non-phytotoxic when used as directed.

USES: Ornamentals, cotton, turf, and nonbearing tree, tobacco, and vine crops, and non-crop areas. Used on a number of crops outside the U.S. and on crops grown for seed.

IMPORTANT PESTS CONTROLLED: Budworms, bollworms, boll weevils, thrips, mites, whiteflies, ants, fleas, ticks, earwigs, sowbugs, crickets, cotton leaf perforator, lygus, loopers, earworms, armyworms, aphids, and others.

RATES: Applied at .025-. 1 lb actual/A.

APPLICATION: Apply when insects appear, and repeat if necessary, usually on a 5-10 day basis. May be used inside greenhouses.

PRECAUTIONS: Do not use in fogging type applicators. Toxic to fish. Buffer the spray solution to a pH of 5-7.

ADDITIONAL INFORMATION: Suppresses spider mite populations. Maintains its activity under high-temperature conditions. May be tank mixed with other products.

ABJ, November, 1989 - Page 717-719

by GLORIA DeGRANDI-HOFFMAN*, DELORES A. LUSBY**, and ERIC H. ERICKSON, JR.*
*Carl Hayden Honey Bee Biology and Insect Biological Control Center
U.S.D.A.-A.R.S., 2000 East Allen Road, Tucson, Arizona 85719
**Rangeland Honey, 3832 Golf Links Road, Tucson, Arizona 85713

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Two serious problems facing the beekeeping industry are the migration of Africanized honey bees into the U. S. and the spread of Varroa mites. Now more than ever beekeepers must manage the genetics of the bees in their colonies if they hope to deal with these problems.

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The strongest tool that a beekeeper has for controlling colony genetics is the grafting needle. Colony characteristics that are favorable to a particular beekeeping operation or are adapted for a specific geographic area can be increased by grafting queens from colonies that possess the desired traits. By grafting their own queens, beekeepers can create lines of bees tailored for the conditions of their apiary sites and beekeeping practices.

A trait that may be an important component in solving Africanized bee and Varroa problems is queen development time. The first queen to emerge destroys the remaining queen cells and becomes the matriarch of the colony. The colony’s behavior and attributes will reflect the genetic composition of the queen and the drones with whom she has mated. Queen development time could be partially responsible for Africanized traits being expressed by bees in geographical areas that previously were inhabited by European strains, if the development period for Africanized queens is shorter than that of European queens. In Africa, queens of Apis mellifera scutellata develop in 14-15 days while European queens require 14-17 days (Anderson, Buys, and Johansmeier 1973). If Africanized queens emerge first the colonies will express many traits associated with that line of bees. Queen development time apparently is an inherited trait. A line of honey bees (hereafter referred to as Lusby bees (LUS) that has been selected for shorter queen development time now has queens with an average development period of 14.1 days (with a range of 12.4-15.8 days). We conducted an experiment to determine the variability in queen development time using a closed population (CP) line of bees composed of stocks that can be purchased from commercial package and queen breeding operations throughout the U.S. (Page and Laidlaw 1982). Larvae from LUS bees were also grafted for comparison. Three CP colonies and two LUS colonies were used for grafting. The resulting queens will hereafter be referred to as CP 1, 2, or 3 or LUS 1 or 2 queens. In this experiment, only 12-24 hour old larvae were grafted (age was determined by size of the larvae). The grafting technique was similar to the procedure outlined by Laidlaw (1981) in which larvae were placed in a drop of royal jelly at the bottom of queen cups. The grafted larvae were then placed in starter-finisher hives containing the same line of bees from which the larvae were grafted. Five days after grafting, the capped cells were placed in individual plastic vials and put in an incubator set at 34.50 degrees C (94 degrees F) and 78% relative humidity (Fig. 1). The incubator was checked every 4-5 hours for newly emerged queens.

The emergence times for CP and LUS queens are shown in Table 1. LUS queens emerged 9.5-10.6 days after grafting (13.5-14.6 days total development time), while CP queens emerged 10.4-11.0 days after grafting (14.4-15.0 days total development time). LUS 2 queens had the shortest average development time. The average development time of LUS 1 queens was not significantly different from those of any of the CP queens.

Table 1. Total development times (egg to adult) of grafted queens from two different strains of honey bees.
Strain	Colony Number	No. of queens	Queen emergence times (days) after grafting
Lusby	1	17	14.6ac
	2	63	13.5b
Closed Population	1	28	15.0a
	2	19	14.4c
	3	23	14.8ac
Means followed by the same letter are not significantly different at the 0.05 level as determined by Scheffe’s S test.

Differences among colonies concerning queen development times are revealed in greater detail by examining the percentage of queens from each colony emerging over time (Fig. 2). Almost 20% of LUS 2 queens had 12-13 day total development times and emerged before other LUS or CP queens. CP 2 had some queens with 13-14 day development times as did both LUS colonies. Most of CP and LUS 1 queens had 14-15 day development times. A relatively small percentage of LUS 1 and CP 2 queens had 15-16 day development times, while almost 60% of CP 1 queens and 20% of CP 3 queens emerged at this time.

Grafting queens and documenting their development time using an incubator is a simple procedure that can be done by any beekeeper. The first step is to determine the range of queen development times in existing stocks, particularly those with other attributes that need to be perpetuated. To do this, graft larvae of the same age. When the cells are sealed, place them in individual plastic or glass vials, and transfer them to an incubator. We use a plastic foam Little Giant poultry incubator, Miller Mfg. Co. Inc., St. Paul, Minn., that costs about $30.00. Check the incubator every 4-6 hours to determine emergence times. Label the vials with the time that the queen emerged, and estimate the total development time. To apply selective pressure for shorter queen development time, introduce only those queens which emerge 9-10 days after grafting. By repeating this process with the off-spring of these queens, the frequency of shorter queen development time can be increased in the next generation. Once this trait is established in a colony, it will be retained even if the colony re-queens itself (assuming that larvae of nearly the same age are selected by the bees to be reared into queens).

Additional studies are currently being conducted using the grafting and selection techniques described here to determine whether queens with shorter development times produce workers with this trait. We are testing factors that could influence development rates. One such factor is temperature which in many insect species strongly influences development rates. In honey bee colonies workers control temperature particularly in the brood nest, and thus may be influencing development rates through temperature regulation.

Shorter development times may be associated with smaller body size. We are examining the size and weight of queens (and possibly workers) with the shortest and longest development times to determine if they differ. If, indeed, queens with shorter development times produce offspring with this trait, they may show resistance to Varroa mite infestations since fewer female Varroa mites will have the opportunity to develop before the adult worker or drone emerges (Camazine 1988).

LITERATURE CITED

Anderson, R. H., B. Buys, and M. F. Johannsmeier. 1973. Beekeeping in South Africa, Dep. of Agric. Technical Services Bull. No. 394.

Camazine, S. 1988. Factors affecting the severity of Varroa jacobsonii infestations on European and Africanized honey bees. In: Africanized honey bees and bee mites. G. R. Needham, R. E. Page Jr., M. Delfinado-Baker, and C. E. Bowman, eds. Ellis Horwood Limited, Chichester, West Suffix, England.

Laidlaw, H. H. Jr., 1981. Contemporary queen rearing. Dadant and Sons, Hamilton, IL.

Page, R. E. Jr., and H. H. Laidlaw. 1982. Closed population honeybee breeding. 1. Population geneties of sex determination. J. Apic. Res. 21: 30-37.

Lusby, Bee Culture - June 1998

The cell size debate may soon be over.

We have been trying to figure out the best brood comb cell-size for some years for our area (Tucson, AZ). It started locally, but then beekeepers in other parts of the United States, and even overseas wanted to know what we thought their best natural brood comb cell size should be also. Eventually we constructed a world map showing thermal zones and corresponding cell size.

To create the map we combined many pieces of information relative to history and world environment, such as: 1) What was the actual recorded cell size prior to the use of artificially enlarged foundation and how was it measured? 2) Where were honey bees capable of living when looking at natural zones of heat and cold during a full year? 3) Were there naturally occurring variables that could change cell size or bee habitat? 4) Would it all fit together - that is, recorded cell sizes compared to the environment they were in?

This map based on atlas composites of hot and cold land area maps accurately reflects the history of recorded cell sizes in published records prior to the use of artificially enlarged foundation. Climate and rainfall were variables allowing habitat transition into and out of recorded zones. Cell sizes are recorded in general for the zones and, where altitude dictates Humboldt’s law, should be used in higher mountainous areas and areas of higher latitude.

Every thermal/cell size zone has a small, medium, and large range to allow for bees to transition into and out of habitat areas as vegetation and rainfall vary throughout the yearly cycle. With smaller cell sizes you gain variability, and with larger cell sizes you have less. We found this published many times.

It seems logical that as you go up the hill bees get bigger to match the colder, higher altitudes, but also, they encounter less habitat to live and breed in. Therefore, when they reach the top of the mountain they have to go down again to regain variability or suffer extinction.

Some of the last of the artificially-enlarged cell size research was done round 1941 with cells of 5.4mm diameter, with the experiments lasting a good four years. Between 1957-1963 other artificially-enlarged cell size research project was undertaken with 5.65 mm diameter cells. The research was considered successful, but the experiments were done in the Romanian mountains. In that area the research showed cell size of natural honeycomb presented great variability, depending upon the altitude. In fact, when all the work was done it was recommended that bee colonies should have at their disposal honeycombs with cells as nearly as possible to the size cell which the bees themselves naturally build.

This made us wonder why we were using “high altitude honey-comb” in “low altitude areas” and then compounding the problem further by placing it into the broodnest. Could this have an impact on the disease and parasite problem? We assumed that since honey bees, mites and other pressures have coexisted for many years, it could be further assumed that something artificial - like oversized cell size - may have disrupted that co-evolved condition, rendering the bees more susceptible to parasites and diseases. So, all we are doing is placing our bees back onto the natural cell size for our area and letting Nature take her course.

We have indeed taken on a puzzle. However, we think we have now found many of the pieces. First, we put our bees on 5.0 mm cell size foundation by making over 40,000 sheets for our brood nests. A long drought forced us to reconsider this cell size though. We have now placed into the field over 4,000 frames of 4.9 mm cell size foundation. That was done in 1997 during the drought and before going into Winter with our bees. We culled heavily to get the job done, shaking down our hives into only 1-2 deep brood boxes to over-Winter.

We spent this past Winter preparing over 10,000 frames with 4.9 mm cell size foundation to put onto our bees between March and the end of July this year. We’ll put on as much extra as we can until frost in November using an additional 10,000 frames of 4.9 mm foundation that will be ready by mid-Summer. That will give us a total of 24,000 frames of smaller, more natural 4.9mm cell size to use once and for all if going back to natural sizing works well for disease and mite control.

The experiments performed in 1941 and 1957-1963 that pointed to the “bigger is better” theory were done with an average of 5-6 colonies. Those experiments changed the world’s beekeeping industry. It became the belief of the day that bigger was better - for more honey, for less swarming, for ease of extracting and spinning out honey. But in the end, what did we gain?

We don’t experiment with only 10 frames or even a few hives as a basis for saying whether or not something works. We’re considered small commercial beekeepers, and averaged 900-1100 hives up until the mites came. We believe well never manage over 700-800 hives with a more intense, smaller, natural bee management system, but we also believe it will be profitable. With El Nino’s help this past year producing lots of plants in the southwest desert, we’ll see how much 4.9 mm foundation our bees can draw out.

We can only come back now as fast as we can draw comb. We’ve been sitting under a mesquite tree now for nearly three years waiting for the chance to draw a large amount of comb. We’re ready. It will be a challenge to see if we can retool all our combs, with the goal of running 700-800 hives, by the end of 1999. all in 4-5 deeps, but if we make it so can everyone else.

Look at the thermal/cell size zone map. So far everything is pretty much matching up. The trouble is, now the questions start. What have we lost by having too-big bees relative to crop pollination? Is there an upper cell size limit for controlling mites relative to altitude and cooler latitudes? What’s the relationship of cell size to disease, chemical contamination, pesticide sprays, inbreeding, pollen and propolis for human cures? Are there limits to taking bees out of one zone and placing them into other zones? If so, is it beneficial, or detrimental?

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Ed and Dee Lusby - deelusbybeekeeper@mailexcel.com
The Lusby’s are commercial beekeepers in Tucson.
You can reach them at 3832 East Golf Links Rd., Tucson. AZ 85713.

ABJ, December, 1997 - Page 837-838

On 11 September Dr. Eric H. Erickson, the director of the Carl Hayden Bee Research Facility in Tucson, Arizona, went with us to two bee locations, in unisolated areas, to test for both tracheal mites and Varroa mites. Samples taken in the center of the brood nest also contained drones where possible. We choose unisolated locations because we wanted to show him, to beat the problem, one must be able to accomplish business as normal in doing bee management within the field. Please note that beekeepers around us have severely lost bees, as we ourselves have, to both mites over the years. When taken, several adjacent yards within 2 miles were being treated, crashing, or being fed to keep them alive. Our bees were building; and at the Carmen yard were very fast drawing new foundation.

We began putting the 4.9 cm cell size in hives in May. We did a second round the end of June and did a third round ending Labor day. The Carmen yard we took samples from was worked Labor Day along with the Knight location. The Carmen yard had been drawing wax and averaged 4-8 or more frames per colony drawn. A few colonies had a full box (10 frames) drawn. The Knight location had less than 3 frames drawn on average and most brood laying was on 5.0 cm comb. Both yards still had 2-3 (3-Carmen 2-Knights) one super hives (nucs) still laying on the larger Duragilt that refused to change. Note these one super hives are now dead, not having survived through to mid-October. So much for Duragilt (5.44 cm).

With smaller 4.9 cm comb which is still bigger than the 4.83 cm comb this country was founded on in Southern latitudes, (Northern latitudes were founded on 4.9 cm to 5.0 cm sizes), we are now getting our Varroa populations down to field tolerant coexistent levels so we can mimic natural environment living conditions. Tracheal mite levels are down there also, having regulated the mite back to external Vagans status, as was the norm condition around 1917 in our country, before we artificially mutated the bee’s thorax and breathing tube bigger on the thorax to create a parasite problem. At 0-6% tracheal mites, bees have no problem coexisting. At 10-11%, Varroa mites are on the cuff for trouble. In Southern latitudes in times of plenty they do fine; in times of dearth the bees do poorly and both require constant management to control secondary diseases. This is on 5.0 cm size comb. At 0-7% varroa mites, changing to 4.9 cm comb sizing, bees draw wax well and hives no longer require constant management to control secondary diseases. Business is back to normal for management in the field. We hope to cut percentages again this coming year 1998 as brood nests are continued with 4.9 cm comb and all frames are converted in our broodnests.

This shows breeding is not all the solution. We figure comb is 1/3, diet is 1/3 and breeding is 1/3. Comb must be put in by half (5) to full boxes to work.

Dee Lusby
Tucson, AZ
Note from author regarding article.

HONEY BEE PARASITES FROM CARMEN
VARROA MITES Colony #	# Bees	# Varroa	# Varroa/100 Bees
A	175	34	19.43
B	186	30	16.13
C	161	39	24.22
D	186	5	2.69
E	157	7	4.46
F	183	13	6.99
G	169	13	7.70
H	148	5	3.38
I	187	2	1.07
J	149	6	4.03
K	185	5	2.70
L	164	7	4.27
M	188	7	3.72
N	156	5	3.21
P	163	8	4.91
Q	179	17	9.50

TRACHEAL MITES IN 30 BEES Colony #	# Tracheal Mites	% Tracheal Mites
A	0	0.00
B	0	0.00
C	0	0.00
D	0	0.00
E	1	3.33
F	0	0.00
G	1	3.33
H	0	0.00
I	1	3.33
J	7	23.33
K	0	0.00
L	1	3.33
M	1	3.33
N	1	3.33
P	0	0.00
Q	2	6.67

HONEY BEE PARASITES FROM KNIGHT
VARROA MITES Colony #	# Bees	# Varroa	# Varroa/100 Bees
A	165	1	0.61
B	186	15	8.06
C	142	13	9.15
D	177	18	10.17
E	168	21	12.50
F	184	23	12.50
G	171	26	15.20
H	186	9	4.84
I	181	53	29.28
J	200	8	4.00
K	189	19	10.05
L	182	4	2.20
M	175	23	13.14

TRACHEAL MITES IN 30 BEES Colony #	# Tracheal Mites	% Tracheal Mites
A	2	6.67
B	3	10.00
C	0	0.00
D	0	0.00
E	0	0.00
F	0	0.00
G	5	16.67
H	8	26.67
I	0	0.00
J	4	13.33
K	2	6.67
L	1	3.33
M	1	3.33