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Working with Nasonia

Published in Resources & Tips. Tags: , , , , .

Nasonia vitripennis is a small parasitic wasp that is involved in a symbiotic relationship with a host organism known as Sarcophaga bullata, or a flesh fly. More specifically, Nasonia is a “parasitoid”, which is a parasite that completely destroys its host. By interrupting the life cycle of Sarcophaga, Nasonia is able to use its host for nutrition and shelter.

I. Sarcophaga Life Cycle

The Sarcophaga host, which is harmless to humans, is associated with animal carcasses. A non-virgin female Sarcophaga will lay a large number of eggs on a carcass. The eggs rapidly develop into larvae (maggots), which will feed on the dead animal tissue. The larvae infest the carcass for nine to ten days before they wander into the soil surrounding the carcass in search of a dry place to pupate. Pupation involves the development of a protective “shell” or casing around the body of the Sarcophaga. Within the casing, a pupa develops into an adult fly in another nine to ten days (figure 1).

As stated earlier, the parasitic Nasonia is capable of interrupting the Sarcophaga life cycle. An adult female Nasonia will lay her eggs within the Sarcophaga pupal casing. This provides a well-protected, nutrient-rich environment for developing Nasonia as they feed on the tender Sarcophaga pupa. The environment within the Sarcophaga pupal casing creates ideal conditions for the completion of the Nasonia life cycle.

II. Nasonia Life Cycle

Nasonia have a relatively short life cycle depending on environmental temperatures and exposure to a light source (figure 2). A female Nasonia lays her eggs in a Sarcophaga pupa by distending a long, thin structure, known as an ovipositor, from her abdomen. The female then deposits thirty to fifty eggs through the ovipositor into the pupal casing. The eggs develop into larvae within one or two days. The larvae then feed on the Sarcophaga pupa as a nutritional source. The larvae will continue to develop over the next eight to nine days, and then pupate, forming a protective casing around their bodies. There are three developmental stages of Nasonia pupae: “white”, “black and white”, and “black” stages, and each stage is more developed than the previous. Nasonia remain in the pupal stage for three to four days. In a process known as eclosion, “black” pupae will break free from their pupal cases as adults These adults eventually emerge by chewing a hole through the host casing.

Figures 1 and 2

III. Benefits of Using Nasonia for Scientific Research

There are many reasons why Nasonia vitripennis is an excellent research organism. For example, its small size, short life cycle, and ability to produce many offspring are advantageous in the study of animal behavior and genetics. Many scientists prefer to work with Nasonia over other research organisms because they are very simple to culture, handle, and maintain.

A. Arresting Development of Nasonia and Sarcophaga

The developmental rates of Nasonia and Sarcophaga are dependent upon temperature and exposure to light. Standard culturing conditions are 25°C and 24 hours of light per day. Cooler temperatures and less exposure to light will slow down the development of both organisms. A temperature of 4°C is ideal for arresting development. By controlling temperature and exposure to light, scientists are able to carefully plan their experiments for maximum efficiency.

B. Observation of Developmental Stages and Timing

The various stages of Nasonia development can be observed at anytime by cracking open a parasitized Sarcophaga pupa. Each stage is an indicator of the amount of time before the Nasonia reach adulthood. This ease of observation makes it convenient for scientists to schedule their laboratories around weekends and holidays, etc.

C. Virgin Isolation and Sex Determination

Virgin Nasonia can be identified while they are still pupae. This allows scientists to easily collect, isolate, and store virgins until needed. Collection and isolation are simple in any of the three pupal stages because the Nasonia are immobile. There are often several distinct differences between male and female Nasonia in the pupal stage (figures 3 and 4). Male pupae are smaller in body size and have short wings (a). Female pupae have a larger body size, long wings that wrap around the abdomen (b), and a visible ovipositor (c).

Figures 3 and 4

Once Nasonia reach adulthood these differences remain except that female wings now extend beyond their abdomen (figure 5). There are also other differences between males and females that become apparent, such as body color, leg color, and type of antennae. Females have a dark-colored body, legs, and antennae. Males have a body with a green sheen, yellowish legs, and light-colored antennae.

Figure 5

IV. Culturing and Handling of Sarcophaga and Nasonia

The basic requirements for culturing and handling the host organism, Sarcophaga, and Nasonia are as follows: Important! The Nasonia and Sarcophaga are both harmless to humans, although there have been some reports of allergic reactions to the hemolymph in the flesh fly pupae. Teachers and students are encouraged to wash their hands before and after handling Nasonia and Sarcophaga.

A. Sarcophaga Hosts

  1. The non-parasitized Sarcophaga hosts are sent in the early pupal stage. Refrigerate the hosts immediately upon arrival. Refrigeration at 4°C will keep the hosts viable for three to four weeks. Important! If unrefrigerated, adult Sarcophaga will emerge in approximately ten days. Keep the hosts refrigerated at all times. This will ensure that they remain as young and tender as possible for parasitization.
  2. At the time of culturing, remove only the number of hosts needed.

B. Nasonia

Note: Nasonia are shipped in the pupal stage while still in the host.

  1. Upon receipt, you may want to check the stage of the Nasonia. To do this, remove a parasitized host from the culture tube. Gently crack open the thin pupal casing of the host which contains the Nasonia pupae (figure 6).
    Figure 6 - Cracking Open Host
  2. Once the Nasonia pupae are exposed, note the stage that they are in. This should be the stage that the rest of the culture is in. Place the cracked open host containing the Nasonia pupae back into the culture tube. Note: The pupae may be at varying stages, so it is important to note the stage that the majority of the pupae are in for planning purposes. If larvae are present, they will not develop after the host has been cracked open. If adults are present they can be used immediately for sub-culturing. Assume that these adults are non-virgins, so do not use them for experiments needing virgins.
  3. If you need virgin Nasonia for your experiments, they should be sexed in the pupal stage (see Section III, Part C: Virgin Isolation and Sex Determination).
  4. Adult Nasonia will be needed for culturing, so you will need to incubate the pupae at room temperature until they become adults. Use the guidelines in table 1 to plan accordingly. Note: Ideally, while incubating, Nasonia should be exposed to cool, indirect light for 24 hours a day. Fluorescent light works well. Light that emits too much heat will harm the Nasonia.Table 1
    Time Consideration: If, at any time, you want to slow down the developmental rate of the pupae to fit into your schedule, you can place the culture tubes in the refrigerator. A refrigeration temperature of 4°C is ideal.
  5. Once they emerge as adults, allow the males and females at least 24 – 48 hours together before sub-culturing. This will ensure that the males and females have had a chance to copulate.
  6. In order to subculture, obtain a clean, empty culture tube.
  7. Obtain the stock culture of Nasonia and tap the tube on the table so the Nasonia fall to the bottom. Remove the cotton plug from the stock and invert the empty culture tube over it. Nasonia will naturally crawl up from the stock culture tube into the empty culture tube. Make sure that the lips of the tubes are lined up so that the Nasonia do not escape.
  8. Allow some Nasonia to enter the empty culture tube. Make sure that females are present because they will be
    the Nasonia that parasitize the hosts (see Section III, Part C: Virgin Isolation and Sex Determination).
    Note: Alternatively, you can temporarily immobilize them by placing the culture tube in the refrigerator for fifteen to twenty
    minutes. This will slow the Nasonia down enough so that you can transfer them by hand to new culture tubes, etc.
  9. Quickly replace the cotton plugs in both tubes.
  10. Tap the new culture tube on the table so the Nasonia fall to the bottom. Remove the cotton plug and add several hosts. Replace the cotton plug.
  11. Label the new culture tube appropriately with the Nasonia strain and the date.
  12. Allow two or three days for the females to lay their eggs. Depending on conditions, the larvae will pupate after 9 –10 days. After this time, a parasitized host can be cracked open, and the Nasonia offspring can be isolated before reaching adulthood (see Section III, Part C: Virgin Isolation and Sex Determination).
  13. Within 14 –15 days after the pupa is parasitized (if incubated at room temperature with 24 hours of light), adult offspring will emerge. Important! If you are working with various strains of Nasonia, make sure that you only work with one strain at a time. Thoroughly clean your work station of one strain before working with other strains.
  14. Dispose of Nasonia as needed (see Section IV, Part D: Disposal).

C. Nutrition

Adult Nasonia can live without food for three or four days, so plan accordingly. If you need to feed the emerged Nasonia, you can add a drop of 4% sugar solution (4 g sucrose/100 ml of water) or honey to the culture tube. For best results, use the Nasonia as soon as they reach adulthood so feeding is not an issue.

D. Disposal of Nasonia and Sarcophaga

Nasonia and Sarcophaga can be disposed of using two methods:

  1. Create a “morgue” by partially filling a container with 10% isopropyl alcohol. Drop the Nasonia and Sarcophaga to be euthanized into the alcohol and cover the container with a lid. OR
  2. Place Nasonia and Sarcophaga to be euthanized into a container and cover it with a lid. Place this container into a freezer and let it sit for at least two hours.
  3. After euthanization, the container used for either method can be disposed of in a wastebasket.

V. Haplodiploid Inheritance in Nasonia

The unique reproductive strategy of Nasonia is known as haplodiploid inheritance and is also common to many other insects, such as bees. Nasonia exhibit a form of asexual reproduction, known as parthenogenesis, in which an organism is able to develop from an unfertilized egg. As a result, when the eggs are not fertilized, offspring will be haploid (n) and male. In turn, when the eggs are fertilized by a male, the chromosome number is restored to the diploid state (2n) and the offspring will be female (figure 7).
Figure 7
Once a female has copulated, she will store sperm until needed. She is able to “choose” when to fertilize her eggs. For example, the offspring from a non-virgin female are usually 95% female and 5% male. This occurs because the female chooses to fertilize 95% of her eggs with the sperm she has stored, resulting in diploid female offspring. The other 5% of her eggs remain unfertilized and therefore develop as haploid males.

VI. Genetics of Nasonia

When performing experimental genetic crosses, be sure to keep careful records of all experiments, giving dates of hosting, results of crosses, and comments about technique.

As an example, a virgin female wild type Nasonia is crossed with a male scarlet eye mutant Nasonia. The dark eye of the wild type is a dominant trait and is represented by “D”. The scarlet eye is recessive and is represented by “d”.

Since the F1 generation is the result of two homozygous strains of Nasonia, the wild type parent has only one possible characteristic in the gametes, wild (D). The scarlet parent also has only one possible type of gamete (d). The F1 generation can therefore be diagrammed as follows:

Note: The male Nasonia is haploid, so the chromosome carries only one allele for the scarlet-eye trait, indicated by the “d” in the diagram. The “–” represents the missing chromosome in the haploid male. The female Nasonia carries two alleles for the dark-eye trait, indicated by “D”. When the male and female alleles combine, they produce dark-eyed females (“Dd”). The female Nasonia is also able to produce dark-eyed male offspring through parthenogenesis, represented by “D–”.
Table 2
Since the dark-eyed trait is dominant, all of the F1 Nasonia will appear to be of the wild type, even though some are not homozygous strains.

A brief survey of methods for working with Nasonia is all that has been possible within the scope of this manual. WARD’S carries a full line of activities using Nasonia, and we urge you to take advantage of these activities, which will provide greater detail and a more in-depth look at the versatile Nasonia.

Note: You can also familiarize yourself with Nasonia by viewing WARD’S Working With Nasonia training video.

This guide is also available in PDF format on wardsci.com.

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Working with Insects

Published in Resources & Tips. Tags: , , , , .

Blowflies and Fleshflies

Calliphora sp., Sacrophaga bullata

In addition to blowflies and fleshflies, the order Diptera also includes mosquitoes, midges, and sawflies. All of these insects provide examples of complete metamorphosis.

You should receive your flies as pupae. When you receive them, place the pupae in a dry dish and put the dish in a screened cage; generally a cage 18″ x 18″ x 18″ will provide enough room for 100 adults. Incubate the pupae at 25°C (77°F) until the adult flies emerge. They should emerge within 5 – 14 days and continue to emerge for three days. When the adults begin to emerge, place one small bowl of sugar and one small bowl of water in the cage. The water should contain a small piece of sponge to prevent the flies from drowning.

Once all the adult flies have emerged, they will deposit eggs. Prepare a medium for the larvae by mixing one part sawdust with three parts dry dog food. Slowly add tap water to make a “mush”. Put this mixture in a pan about 11⁄2 –2″ deep and place a slice of scored raw beef liver on top. Once the medium has been prepared, place the pan in the adult cage. Add water to the medium daily to keep it damp, but not wet. In several days white larvae should be visible on the liver, especially the underside.

Figure 1
After about one week, remove the medium pan with the larvae. Line the bottom of a larger container with paper towels. A plastic dishpan works well. On top of the paper towels add a jar lid for a spacer, then place the medium pan with the larvae on top of this and cover the container with a screen (Figure 1). Continue to add raw liver as needed. In approximately six days the larvae will migrate over the sides of the medium pan and fall onto the paper towels, where they will develop into brown pupae in about four days. These pupae may be incubated immediately to produce a second adult generation or incubated at 4°C (39°F) for up to eight weeks.

Butterflies and Moths

Lepidoptera

Lepidoptera are homometabolous insects like those from the order Diptera and have a life cycle that demonstrates complete metamorphosis.

You will be supplied with pupae when you order from WARD’S; however, it is also possible to collect adults from nature and induce the gravid females to oviposit or just collect the eggs in the field. If you do collect your own specimens, be sure to note the kind of plant where the animal or eggs were found. You will need to make sure that you have the appropriate fresh plant material for the larvae to eat when the eggs hatch.

Keep the eggs in a Petri dish until they hatch. After the larvae hatch, they should be transferred to a larger container, and, as they grow, they should be moved to increasingly larger containers to prevent overcrowding. The bottom of the container should be covered with a substrate of loose soil for moths that pupate below ground. Sticks should also be placed in the container for other species that pupate above ground and for the adults to sit on as they dry their wings. Clean the container as needed to prevent the buildup of frasse (caterpillar excrement). Fresh plant material should be provided daily. It is important to choose plant material specific for the species. Reference books can help you determine the appropriate plants to choose for each species.

Cockroaches

Periplaneta americana

The cockroach is from the order Blattari and is an excellent example of simple metamorphosis. It is also widely used in experiments on physiology and testing insecticides.

Cockroaches are best reared in an aquarium or a terrarium that is covered with a tight-fitting screen. The container should be kept at room temperature in the dark or in a darkened area of the room. Stack several boards separated by 1⁄4 – 1⁄2″ spacers, to make a series of “apartments” where the roaches can congregate. Place a band of petroleum jelly around the top of the container to prevent the roaches from climbing to the top. Replace the petroleum jelly occasionally. Cleaning is rarely necessary and should be done only as necessary.

Keep the roaches supplied with water (we recommend WARD’S insect watering device, 14 W 7510) and food (dog biscuits work well) at all times. Supplement the dog biscuits with pieces of potatoes, apples, lettuce, and stale bread on a weekly basis.

Under these conditions, the colony will reproduce and survive indefinitely. Other cockroach species, such as our giant hissing cockroach may also be cultured in a similar manner. See the specific culture instructions included with each culture for more information.

Confused Flour Beetles

Tribolium confusum

The confused flour beetle is a from the order Coleoptera and is from the family Tenebrionidae. It is mainly used to demonstrate complete metamorphosis. T. confusum can be a serious pest of stored food products so you should take the necessary precautions to prevent their escape.

Place the beetles in glass jars or culture dishes and add whole wheat flour, white flour, or cornmeal as a culture medium. You may also use WARD’S Tribolium medium. It is not necessary to add water to any of the chosen media.

Transfer the beetles to a different container with fresh medium periodically to avoid overcrowding. When transferring the beetles, keep in mind that they emit a disagreeable odor when disturbed, so using an aspirator is not recommended. Instead, transfer the beetles with forceps or a camel’s hair brush.

Tribolium beetles take about six weeks to develop from egg to adult at 27°C (81°F) and 40% humidity. Adults generally live between six and twelve months, but may live as long as three years.

Crickets

Gryllus, Acheta

Crickets, along with grasshoppers and katydids, are from the order Orthoptera and exhibit simple metamorphosis.

Crickets are easy to raise, they can simply be placed in an aquarium or glass jar. If the container is higher than eight or nine inches a cover is not necessary, although shallower containers should be covered to prevent escape. A single pair of crickets require only a 1 L jar, but larger accommodations should be supplied for colonies.

Cover the bottom of the container with approximately 1″ of damp sand. The damp sand will provide enough water for small crickets, but adults and larger juvenile crickets should have a small watch glass or similar container filled with water as well. Add a sponge to the water container to prevent drowning. You may wish to keep the smaller, immature crickets in a separate container, since they may still drown in the water container.

Crickets should be given ground rolled oat paste made by adding a small amount of sugar, skim milk powder, and water. Spread the paste on sheets of paper and allow it to dry. Cut the paper into 1″ squares and place one in the aquarium or jar every two or three days. They will also eat lettuce, grass, fruit, and almost any food that does not mold quickly. You may also use WARD’S cricket food.

Dermestid Beetles

Dermestidae

Most species in the order Coleoptera and family Dermestidae are scavengers that feed on a variety of organic matter. Often used for a variety of biological experiments, dermestid beetles are also used in skeleton preparation. The larvae will clean flesh and cartilage from a dried carcass, leaving the bones ready to be rinsed, degreased, bleached, and assembled. Despite their beneficial role as scavengers, the larvae can also be serious pests, destroying textiles, stored food, and museum collections. The adults feed on flowers and seldom cause damage, but it is still recommended to take precautions to prevent these beetles from escaping.

Place the dermestids in any container that can be sealed. Add dried protein such as dried meat, cereal, insects, or fish food as a culturing medium. We also recommend WARD’S dermestid medium. While atmospheric humidity generally provides enough moisture for the beetles, you may also add a small piece of cotton or filter paper that has been dampened with water to the container. Clean the container by filtering the debris through a sieve that will retain the larvae.

The beetles will produce two or three generations in a year. You may start a new culture by transferring a portion of an existing culture to another culture container with fresh media or by transferring newly emerged adults to a new container for mating and egg production.

Dragonflies and Damselflies

Aeshnidae, Coenagrionidae

These common predaceous insects are members of the order Odonata and they exhibit simple metamorphosis. Immature stages are aquatic and do not resemble the adults as most insects do when developing through simple metamorphosis. The aquatic stages are called naiads, which are also voracious predators.

Dragonfly and damselfly naiads can be collected almost any time of the year and maintained indoors over the winter. Some species may remain naiads for several years before becoming adults. Adults will emerge in the spring.

The naiads can be kept in a small aquarium and require live food. Smaller naiads will feed on Daphnia and similar small crustaceans, while larger forms will eat other aquatic insects, mosquito larvae, and even small fish. An ideal culture setup for these aquatic predators is a five to ten gallon tank with plants such as Elodea and Vallisneria. At least one floating plant should be included to enable the insects to emerge from the water as they transform into adults. Add some tadpoles and pond snails, then introduce the dragonfly and damselfly naiads. Daphnia and mosquito larvae should be added every two or three days as needed for food. The aquarium should be covered with netting to prevent the adults from escaping into the classroom. Once the naiads have all emerged, you may capture the adults and release them outdoors.

Mealworms

Tenebrio

Mealworms are from the order Coleoptera and the family Tenebrionidae and are ideal as food for insectivorous animals, as well as subjects for various lab experiments.

Mealworms produce a single brood laying eggs from May through late October and producing larvae from fall through the following spring. A culture started in April will usually produce enough mealworms for use in experiments in the following school year.

Mealworms can be raised in containers such as a wooden box, a glass jar, an aquarium, or any other similar container. Place wheat bran or a similar wheat meal product in the bottom of the container to a depth of several inches. The wheat meal may be mixed with chick mash or hog meal. For food that has already been prepared, we recommend WARD’S Tenebrio nutrient. Cover the meal with a damp, but not soaked, piece of burlap or paper towel, and then place an additional 1– 2″ of food on top of the burlap. Add the mealworms and cover the container with a screen to keep the adult mealworms from escaping. Keep the container in a relatively dark corner of the room at an average temperature. Add slice of raw potato weekly to maintain the culture.

Milkweed Bugs

Oncopeltus fasciatus

Milkweed bugs are from the order Hemiptera and the family Lygaeidae. They can be used to illustrate simple metamorphosis and the morphology of a typical hemipteran.

O. fasciatus are clean and thus are easily maintained. First, establish a colony in Petri dishes or glass culture dishes, then keep the colony in an aquarium. Line the container with paper towels to absorb excreta. Place a small ball of cotton or cheesecloth in the container to collect eggs. Eggs will be deposited in groups of ten to fifteen and as they incubate they will change color from yellow to deep orange. Because the adults and nymphs will feed on the eggs, the cotton containing the eggs should be removed daily.

Milkweed bugs feed on various species of milkweed, but prefer dried milkweed seeds. These can be scattered on the bottom of the container. Also provide the insects with water in a small vial with a cotton plug wick. Change the water and cotton plug every fourth day.

O. fasciatus develop from egg to maturity in approximately six to seven weeks. See the table for the time required for various stages of development. The markings on milkweed bugs make it easy to determine the sex. A female will have two black spots on the second abdominal segment, a band on the third segment, and two black spots on the fourth segment. The male, on the other hand, lacks the spots on the second segment, while the third and fourth segments have a solid black band instead of spots.
Figure 2

Mosquitoes

Culex, Anopheles

Mosquitoes are from the order Diptera and, as such, develop through complete metamorphosis. However, these insects have aquatic larvae unlike the flies from the order. Mosquitoes are often used in medical studies, so there is an abundance of literature detailing various methods of rearing and maintaining mosquitoes in laboratories. The method detailed here should suffice for normal development of both Culex and Anopheles.

Begin a colony by collecting egg masses or larvae. Handle gravid females with extreme care. Place the eggs inside a cork ring and put the cork in a glass jar filled with water. The ring keeps the eggs from adhering to the sides of the jar.

The most convenient medium for rearing larvae is a wheat infusion. You can make a wheat infusion by boiling wheat grains for about five minutes. Add 250 grains of boiled wheat to 2 L of water in an enamel pan or glass jar. Allow bacteria to grow for approximately two or three days, then inoculate the culture with material from an older culture that supports a thriving population of flagellates and ciliates. To maintain an alkaline pH, add 1 g of calcium carbonate. This culture should be usable for two or three weeks. After another five to seven days, add the mosquito larvae. Supplement the wheat infusion diet with yeast by adding a small pinch daily. If scum develops on the surface, reduce the amount of yeast added, subdivide the culture, or move the smaller mosquito larvae to another culture.

When the larvae transform into pupae, remove them from the culture with a pipet and place them in a pan of clean water. Spread chaff, broken cork, or other similar material on the surface to keep the pupae separated and to provide the adults with a surface to emerge from the water. Change the water daily to prevent the formation of scum on the surface. Put the pan in a cage large enough to provide enough space for the adults to fly. To capture the adults, cover the perched mosquitoes with a vial.

Breeding adult mosquitoes is not a recommended procedure for the classroom because of the space and blood meal requirements.

Termites

Zootermopsis

Termites are relatively primitive insects from the order Isoptera that develop through simple metamorphosis. Termites from the genus Zootermopsis are native to the West Coast of the United States and are the host organisms for many protozoa such as Trichonympha, Trichomonas, Streblomastix, Hexamastix. The majority of these protozoa have a mutual relationship with the termite where each organism benefits from the relationship. In this particular relationship, the protozoa digest wood cellulose in order for the termites to absorb the nutrients.

Zootermopsis require relatively high humidity at ordinary room temperature, but can survive in a wide range of temperatures, although ideally temperatures should not exceed 20°C (68°F). Colonies should be maintained in a large culture dish with rotten wood as the culture medium. Keep the wood moist by adding a few drops of water every two or three days. Fungi are a necessary part of the diet, but be sure to prevent the overgrowth of mold. Only wood that does not appear to be heavily infected with fungus should be used for food. Cultures should also be kept in a dark area.

The culture contains nymphs and adults from two castes. Soldier termites have large heads and strong mandibles and are readily distinguished from the smaller termites that have only reproductive functions. Zootermopsis do not have a worker caste, since the nymphs perform the function of cleaning the nest. Some of the nymphs will develop into secondary reproductive forms, which are recognizable by their light brown color. To establish a permanent colony, separate these nymphs from the adults. Remove any diseased termites immediately.

This guide is also available in PDF format on wardsci.com.

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Working with Hydra

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Introduction

Hydra are freshwater coelenterates found throughout the world. They range in size from less than 1 mm to 5 mm. Their simple structure consists of a polyp— a slender stalk with a row of tentacles surrounding the mouth at the top and a pedal disc at the base, by which the Hydra attach themselves to substrate in streams and ponds. Voracious predators, Hydra’s tentacles are armed with stinging cells (nematocysts) that “harpoon” their prey (usually small crustaceans). The tentacle can then rapidly retract to draw the food into the mouth.

Common species include Hydra vulgaris (brown Hydra) and Chlorohydra viridissima (green Hydra). Green Hydra differ from brown Hydra in that their green color is caused by their symbiont, an alga (Chlorella) Green Hydra are also smaller than brown Hydra, ranging from about 0.5 to 2 cm.

Tissue Function

Hydra’s two main cell layers, the epidermis (outermost cell layer) and gastrodermis (inner cell layer) are separated by a thin mesogleal plate, an “acellular”substance. These cell layers are incipient tissues, consisting chiefly of one cell type but containing a number of other cells; they have broad functions rather than the narrow functions typical of true tissues.

The epidermis consists of vacuolated musculo-epithelial cells containing muscle threads (myonemes). These muscle threads attach to the mesogleal plate; when they are contracted, the Hydra’s body shortens. Epidermis gland cells on the pedal disc form adhesive secretions. Small interstitial cells are packed in at the base of the musculo-epithelial cells; they are especially prominent in the “growth zone”of the anterior part of the column region and at points where buds or sex organs develop. These cells are unspecialized, and can replace cells of any other type. Many interstitial cells become nematocysts (stinging cells).

Large nutritive-muscular cells in the gastrodermis feature transverse myonemes that lengthen the body when they contract. The gastrodermis is specialized by region, as shown by the distribution of cells and the changes in nutritivemuscular cell form. Mucous gland cells are abundant near the Hydra’s mouth, and enzymatic gland cells secrete enzymes for extracellular food digestion. The column contains gastrodermis where digestion and absorption occurs; the stalk region has a low, inactive gastrodermis.

Reproduction

Hydra usually reproduce asexually, by budding. This process lasts two to four days, ending with the daughter Hydra detaching from the parent to become a separate organism. Hydra can also reproduce sexually, although this is much less common. Differentiation must be induced by environmental factors such as high carbon dioxide levels or a change in temperature; sexual reproduction usually occurs in the autumn.

Different species of Hydra are either dioecious or hermaphroditic. Testes form from interstitial cells that produce a swelling in the upper third of the body. Spermatogenesis results in flagellated spermatozoa. In the ovary, a large, yolkfilled oocyte forms in a manner similar to the testes. After the epidermis ruptures over the mature ovum, it is fertilized by spermatozoa released into the surrounding water. The fertilized ovum remains on the parent’s body, where it undergoes cleavage and gastrulation to become an embryo. A protective shell, or theca, forms around the embryo, after which it detaches from the parent. The egg may remain dormant for some time before the embryo emerges as a small immature polyp with a mouth, body, and tentacles.

Care and Feeding

Hydra are best kept in an aerated aquarium or tub. Keep the temperature relatively low (18 – 21°C), provide a steady source of food, and ensure the water supply is free of contaminating chemicals. Brown Hydra prefer colder temperatures and tend to be larger and healthier in the winter months.

Note: Hydra cultures, under usual methods of cultivation, undergo a period of depression in which the animals refuse to feed, the tentacles fail to expand, disintegration sets in, and the colony dies out. Therefore, it is recommended that you cultivate more than one culture at a time.

Use only pond water or a mixture of pond water and deionized water. If tap water is used, remove salts with a water conditioner such as WARD’S Water Conditioner; the conditioned water can be used immediately. If the water is filtered, it is not necessary to change the water; however, if it is not filtered, the water should be changed daily.

Hydra may be fed the larvae of brine shrimp (larger hydra may also feed on Daphnia). Brine shrimp are easy to raise and maintain in the lab: Fill a brine shrimp hatchery with one liter of salt water and two level teaspoons of brine shrimp eggs. Aerate the hatchery and allow eggs to hatch; this will take approximately two days. Collect the larvae by stopping the aeration and placing a light at one end of the hatchery. The brine shrimp will be attracted to the light, separating from the eggs that failed to hatch. Remove the larvae with a pipet and place them in an aquarium net lined with a piece of cloth. Rinse the brine shrimp under treated tap water to remove the salt, then pour them into a container of treated tap water. Use the brine shrimp immediately, as they will die quickly in tap water.

Feed the Hydra daily by scattering the brine shrimp over the colony with a pipet. Allow approximately half an hour to an hour for feeding, then pour off the water with the remaining brine shrimp. You may also pour the water directly into a bowl and swirl the water to collect the Hydra; the Hydra will be moved toward the center of the bowl. Return the Hydra to their original container or place them in a new container.

Because green Hydra’s symbiotic alga is photosynthetic, the green Hydra can be sustained for several weeks without food, provided there is adequate sunlight or wide-spectrum artificial light. Keep the temperature below 25°C (higher temperatures promote rapid growth of algae, which will choke out the Hydra). To keep green Hydra healthy however, they should be fed small crustaceans.

Green and brown Hydra can be kept in a refrigerator in jars for two to three weeks without feeding or water changes. After a few days without feeding, the Hydra will begin to rise to the surface for easy collecting with a pipet.

Culturing Hydra

In a proper environment, Hydra will bud profusely and at times produce sexual individuals. Cultures that are fed and cleaned daily will produce spontaneous sexual differentiation.

Note: Aeration will inhibit the development of sexual individuals; if you want to induce development of sexual stages, aeration should not be used.

Temperature reduction will also usually bring about the production of sexual forms. The simplest method is to place the Hydra culture on the bottom shelf of a refrigerator and feed daily.

Hydra Longitudinal Section

Hydra images

This guide is also available in PDF format on wardsci.com.

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Working with Fungi

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Introduction

The Fungi Kingdom includes spore-forming eukaryotes that lack undulipodia (amastigote) at all stages of their life cycle, but some taxonomists include chytrids and oomycetes, which are undulipodia, in the Kingdom.

Almost all fungi are aerobic, and all fungi are heterotrophs, which means that they absorb their food without ingesting it. Fungi secrete powerful digestive enzymes that break down the food material, usually decaying plant or animal matter, into compounds that can be absorbed through the fungal membrane.

Fungi are abundant in most terrestrial habitats. There are about 100,000 known species, a few of which are marine. Because of their abundance and diversity, there are likely many more species to be discovered and described.

Reproduction in fungi is generally asexual and occurs through vegetative spores (conidia). These spores are distributed by wind and water, and are highly resistant to adverse environmental conditions. Under favorable conditions they will germinate and initiate the growth of a new organism. The germinating spore produces a thin, tubular structure called a hypha. Cross walls called septa divide the hypha into “cells”, though the septa rarely completely enclose an entire portion of the hypha. Some hyphae lack septa entirely. Hyphae grow in large masses called mycelia, which constitutes the vegetative body of the fungus. The cell walls of most fungi are composed of chitin, similar to that found in insects, which is very impervious to desiccation, thus enabling the fungi to survive under harsh conditions (fig. 1).

Figure 1

Sexual reproduction can also occur in fungi. This occurs when hyphae of opposite mating types grow together and fuse. The haploid hyphal nuclei grow and undergo subsequent division, but they remain in pairs, one nucleus from each of the two parental hyphae. Such hyphae are called dikaryotic. Nuclear fusion occurs in time, forming diploid zygotes. The zygote then immediately undergoes meiosis to form haploid spores that are distributed in the same manner as the vegetative spores.

Sexual reproduction modes and structures form the basis for classifying the major groups of fungi. There are four phyla of fungi.

Figure 2

1.Zygomycota. These are the “algae-like” fungi. There are no septa in the hyphae, though the reproductive structures are separated from the rest of the mycelium. Genetic material is exchanged in thick-walled zygospores, which are formed by the conjugation of opposite mating types, and then the haploid spores are released from the zygospores. Asexual reproduction is by conidia, resistant spores that develop within the sporangium. The common bread molds are examples. Some examples are common bread molds and parasites of protists, nematodes, insects, and small animals (fig. 2).

Figure 3

2. Ascomycota. The mycelium of Ascomycetes form a cottony mass of multi- branched hyphae. This phylum is characterized by sac-like reproductive structures called asci, which result from the conjugation of two compatible mating strains. Each ascus typically contains eight ascospores, which form when the short-lived diploid nuclei undergo meiosis. Many serious plant pathogens are in this phylum, including Dutch Elm Disease and apple scab. The group also includes beneficial yeasts (vital to baking and brewing industries) and highly prized edible fungi, including the morels and truffles (fig. 3).

3. Basidiomycota. The phylum is distinguished by its reproductive structure, the club-like basidium, which contains the products of sexual reproduction called basidiospores. Germinating basidiospores produce a mycelium that develops septa as it grows. Hyphae of compatible mating types conjugate to form a secondary mycelium. A tertiary mycelium develops to form the familiar reproductive structures of mushrooms, puffballs, and bracket fungi. Some basidiomycetes, the rusts and smuts, are serious crop pests, while others can be used for food or medical products. Some members of this phyla can also be deadly because of their poison. (fig. 4)

4. Deuteromycota. These fungi are often termed “fungi imperfecti” because they lack structures for sexual reproduction. Only asexual reproduction is known to occur naturally, although some genetic recombination has occurred under laboratory conditions. Germinating spores produce mycelia with septate hyphae. The phylum includes economically important species like the pathogenic Candida, as well as Penicillium, source of the powerful antibiotic Penicillin. It also includes an interesting genus of predaceous fungi, Arthrobotrys, which actively captures and feeds upon nematodes. (fig. 5)

Figures 4 and 5

Biotechnology has enabled us to use the unique biochemical makeup of these organisms to our advantage. Fungi are important in the processing of many foods, including tea, coffee, cheese, vinegar, beverages, and breads. They are also influential in the manufacture of industrial chemicals such as acetone, alcohols, and citric acid, as well as in the production of antibiotics and vitamins. Last, but by no means least, is the recycling of organic matter. Fungi are vitally important in initiating the breakdown of matter through decay, helping to return vital nutrients to the soil.

Why WARD’S is Better

  • We maintain all cultures in our own laboratory
  • Cultures are maintained under optimum conditions by subculturing regularly
  • You are assured of getting fresh material in the maximum growth phase
  • Cultures are available in a number of formats to meet specific teaching requirements
    • Tubes
    • Plates (two week lead time may be required)
    • Freeze-dried
    • Jars
  • Fully labeled cultures indicate species, media, and incubation temperature
  • Instructional literature is supplied with every order

Care of cultures

Unpack your shipment carefully. Read the labels on the culture containers for information on storage temperatures and conditions. Most fungi cultures can be kept at room temperature, where they will last for several weeks if kept in subdued light. Avoid excessively high temperatures and keep out of direct sunlight. Loosen the caps on tube cultures – fungi are aerobic. Lyophilized (freeze-dried) cultures can be stored at room temperature if you intend to use them within a week. Refrigeration or freezing is recommended for long-term storage.

Handling Cultures

Precautions: Sterile technique is essential in handling all cultures of fungi. This will eliminate any risk of contamination in the culture, and, more importantly, will protect you and your students from accidental exposures to the organisms. Even though most fungi are non-pathogenic, they should be treated as potentially pathogenic to eliminate all possible risks of exposure.

  1. All media and glassware should be sterilized prior to use. An autoclave or pressure cooker at 15 psi for 15 minutes at 100°C or dry heat sterilization of glassware at 160°C for two hours will suffice. WARD’S prepared media has already been sterilized for your convenience.
  2. Make sure that the work area is free of clutter. Wash the work surface with a strong disinfectant, such as 5% Lysol or 70% alcohol, both before and after use.
  3. Wear a lab coat, smock, or apron to protect your clothes and reduce further the chance of contaminating the cultures.
  4. Avoid any hand-to-mouth operations such as eating, drinking, smoking, or licking adhesive labels while in the lab.
  5. Wash hands thoroughly with soap and water both before and after working with cultures.

Culture Transfers

Materials needed for slants: Inoculating loop or needle, Bunsen burner or similar flame source (CAUTION – do not use alcohol near an open flame), culture, second container with media for culture transfer.

Slants: Follow the sequence of drawings Fig. 6a-e for a successful transfer

  1. Hold both tubes in the left hand (fig. 6a)
  2. Hold the inoculating loop or needle in your right hand. Pass the entire length of the wire through the flame until it all has been red hot (fig. 6b)
  3. While still holding the needle, quickly remove the caps (or plugs) from the tubes, holding them between the fingers of the right hand (fig. 6c).
  4. Flame the mouths of both culture tubes by passing them two or three times through the flame (fig. 6d). Hold the
    tubes almost parallel to the work surface, to reduce the possibility of air-borne contaminants.
  5. Touch the needle or loop to the medium in the culture tube, to be sure it is cooled, then to the culture mass, making sure that a portion of the fungal culture adheres to the loop or needle (you do not need a large amount) (fig. 6e).
  6. Quickly withdraw the needle or loop and insert it into the second culture tube, gently wiping it across the surface of the agar.
  7. Flame the mouths of each tube as before and replace the caps or plugs.
  8. Flame the inoculating needle or loop until it is red hot. You have now completed the tube transfer.

Figure 6

Materials needed for plates: Sterile Petri plates, bottled media, large beaker, hot plate, thermometer, inoculating needle or loop, Bunsen or similar flame source

Plates: Use sterile technique as outlined above.

To make plates using bottled solid media:

  1. Place the bottle of media in a large beaker. Add water to the beaker until the water level is just above the level of the medium in the bottle.
  2. Loosen the cap on the bottle.
  3. Put a thermometer in the beaker; set it on a hot plate and heat the water to boiling.
  4. Meanwhile, set the Petri plates (each bottle of media will supply 5 – 7 plates, depending on the thickness poured) on the work surface
  5. Boil the water gently for several minutes until the medium is completely liquefied. Swirl the bottle gently to be
    sure all is melted.
  6. Turn off the heat. Allow the water temperature to cool to between 45°C and 50°C.
  7. Using an insulated glove, pick up the bottle, remove the cap, and flame the bottle mouth.
  8. Lift the lid of a Petri plate just enough to admit the neck of the bottle, and pour the first plate, using just enough medium to slightly more than cover the bottom of the plate. Replace the lid.
  9. Swirl the plate gently to distribute the medium.
  10. Flame the mouth of the bottle and proceed to pour the rest of the plates

To transfer cultures to plates:

  1. Follow aseptic technique as described for slants
  2. Lift the lid of the Petri dish just enough to introduce the inoculating loop. Gently wipe the needle or loop across the culture (you only need a small amount ).
  3. Replace the lid, and then lift the lid of the sterile petri dish containing fresh medium just enough to introduce the needle or loop. Gently wipe the needle or loop across the surface of the media, then replace the lid and sterilize the needle or loop as before. You have now transferred the fungal culture to the plate.

Lyophilized (Freeze-dried) Cultures: Specimens to be lyophilized are grown under optimal conditions to achieve the maximum growth rate. 0.5 ml cell suspensions are then taken and pipetted into sterile vials and freeze-dried. Instructions for rehydrating freeze-dried cultures are included with each culture that we ship. The great advantage of lyophilized cultures is their longevity. Unopened cultures can be stored under refrigeration for ten years or more. Cultures stored at room temperature can be kept for up to two years. Most media supplied with lyophilized cultures can be stored for up to six months. Lyophilized cultures can be conveniently stored, so that you will have them on hand whenever you need them. There is no need for specific ship dates to meet your schedule. Cultures are easily rehydrated with fungi outgrowth occuring 7 – 14 days.

Macrofungi Cultures: Corprinus and Schizophyllum cultures are shipped in jars. They can be subcultured, using sterile technique, by excising a bit of the media upon which they are growing, and transferring it to fresh medium in another jar. The jar should be large enough to allow the reproductive structures to develop. Other larger fungi may be kept in jars to display the morphology of the reproductive structures and in some cases part of the mycelium. Larger fungi may also be maintained in a terrarium. They should be collected with a large portion of the substrate to be sure that the mycelium is included. Mushrooms, bracket fungi, coral fungi, and others are easily collected and can demonstrate diversity within the fungi kingdom. They can also be used to illustrate the structural differences between the phyla.

Culture Disposal/Handling Spills: Safety concerns should always be kept in mind when conducting any microbiology work. You should have on hand at all times:

  • Autoclavable Bio-Hazard bags
  • Paper towels
  • Latex or vinyl gloves
  • Beaker tongs or bottle forceps
  • 70% alcohol or 10% Lysol in squeeze bottles

To dispose of cultures:

  1. All cultures must be autoclaved at 121°C at 15 psi for 15 minutes
  2. Contents of containers can then be discarded. If autoclave is unavailable, soak in bleach or incinerate.
  3. Glassware should then be washed in hot water and rinsed well for re-use.
  4. Note: Do not pour melted agar down the drain, as it will solidify and plug the drain.

To handle a spill:

  1. Put on protective gloves. Pour disinfectant (70% alcohol or 10% Lysol) on all broken glass and contaminated surfaces. Extend coverage at least 3” beyond contaminated area. Make sure that all ignition sources are eliminated if using alcohol.
  2. Cover the spill area with paper towels. Add more disinfectant solution to saturate toweling. Allow to stand for 30
    minutes.
  3. Using gloves and tongs or forceps, pick up all broken glass, residue, and saturated paper towels and place in the Bio-Hazard bag.
  4. Disinfect the area once more as in steps 1– 3. Dispose of your gloves in the Bio-Hazard Bag.
  5. Seal the Bio-Hazard bag and autoclave contents for disposal.
  6. Wash hands thoroughly.

Special Techniques

General: The techniques previously outlined apply to most fungal cultures. Some special techniques apply to specific fungi:

Saprolegnia ferax, (water mold). This species is best grown on cornmeal agar. It can be grown in liquid medium by adding sterile rice grains to distilled water and inoculating it with Saprolegnia.

Demonstration of sexual reproduction in fungi can easily be done using Rhizopus stolonifer, Phycomyces blakesleeanus, or Mucor hiemalis.

Inoculate plus and minus strains on opposite sides of a Petri plate containing sabouraud dextrose agar or potato dextrose agar. Be careful not to mix the strains when plating them.

When hyphae of the opposite strains grow to meet in the center of the Petri plate, a line of mature zygospores will develop where the strains meet. Have students observe these under a dissecting microscope and sketch what they see.

Making wet mount preparations:

Observe the mold colony under a dissecting microscope. Following sterile technique, use an inoculating needle or loop to remove a small amount of mycelium bearing conidia or sporangia. Place this in a drop of water on a clean microscope slide. Tease it apart with the needle, if necessary. If staining is desired, add one or two drops of methylene blue. Cover with a coverslip and observe under low (40X) then high (400X) power of a compound scope. Draw what you see.

Constructing a moist chamber:

Use WARD’S Silicone Culture Gum to construct a moist chamber, so that you can grow a fungus as a slide culture and observe the entire life cycle. This simple yet beautiful preparation will let you see all phases in the growth of the fungus. The fungus is grown on a block of agar under a coverslip on a glass slide.

  1. Roll a marble-sized ball of culture gum and flatten it to form a disc about 1⁄4” thick.
  2. Press the disc to a clean microscope slide. Use a cork borer to cut and remove the center of the disc.
  3. Immerse the slide into 70% alcohol to sterilize and let dry. Cover with a sterile coverslip.
  4. Use a sterile scalpel to cut a small block of agar. Remove the coverslip from the slide.
  5. Place the agar block in the center of the slide. Using a sterile needle, inoculate the center of each of the four sides of the agar block with mycelia or spores. Replace the coverslip. You can observe the growth of the fungus over a number of days. If necessary, remove the coverslip and add a drop of distilled water to keep the chamber moist.

This guide is also available in PDF format on wardsci.com.

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WARD’S offers a wide variety of fungi cultures, kits, and lab activities.

  • Predatory Fungi Kit - For a dramatic demonstration of predatory behavior, add a nematode to the culture of fungus you grow. The fungus, commonly found in soil, has adhesive loops at the ends of its hyphal branches. As the nematode passes through a loop, it becomes trapped, and the fungus digests it.
  • Introduction to Genetics: A Dihybrid Cross in Yeast Lab ActivityIntroduction to Genetics: A Monohybrid Cross in Yeast Lab Activity - Students will investigate the concepts of inheritance, dominant and recessive alleles, and phenotype versus genotype, as well as learn how to predict phenotypic ratios. Using yeast allows students to obtain results much more quickly than with other organisms, and the results are observable with the naked eye.
  • Introduction to Genetics: A Dihybrid Cross in Yeast Lab Activity - Dihybrid crosses can be complex and abstract in nature, but this activity allows students to perform a cross that clearly demonstrates the properties of dominant and recessive alleles, genotypic and phenotypic ratios, independent assortment, and F1 and F2 inheritance patterns on just two Petri plates.
  • Introduction to the Yeast Life Cycle Lab Activity - Using select strains of Saccharomyces cerevisiae (common Baker’s yeast), students can observe the entire yeast life cycle, including sexual and asexual reproduction, diploid and haploid life stages, and sporulation.

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Working with Drosophila

Published in Resources & Tips. Tags: , , , , .

Drosophila melanogaster is an excellent organism for use in the study of genetics. Its small size, relative ease of maintenance, short life cycle, and its ability to produce many offspring have lent it to the demonstration of Mendelian inheritance in every location from the elementary school to the university laboratory. From the thousands of available strains, WARD’S has selected an assortment of over forty types, which serve most purposes.

The basic requirements for growing and working with Drosophila are as follows:

Media

Many different types of media have been used for the culture of Drosophila. The simplest is just a slice of banana. However, each medium has its drawbacks. In order for a medium to be useful, it must be solid and dry enough so the adult flies do not drown, and it must inhibit the growth of molds, which are almost always present on the flies and in the air.

An improved dry instant Drosophila medium, developed at WARD’S, is recommended for general culture work. Students may mix their own as needed and no sterilizing is necessary. The instant medium is available in containers of 1 L, 4 L, or case lots of six 4 L packages. The medium is complete with dry yeast, measuring cup, and instructions. Use a measured quantity of dry instant medium and place it into a culture container. Then add an equal quantity of tap water without mixing. The water is rapidly absorbed and the medium is ready for flies to be introduced in less than one minute. WARD’S uses this medium to ship all Drosophila cultures.

Handling Flies

Materials needed for working with flies are: a Petri dish, an ice pack or cryolizer, a magnifying device (binocular dissecting microscope, large magnifying glass, etc.), a light (goose-neck desk lamp, a microscope lamp, etc.), a camel’s hair brush, a white viewing surface, and a “morgue” bottle (any bottle with a cap, partially filled with alcohol, mineral oil, etc. into which dead flies and those to be killed can be dropped).

Put the culture of flies in a refrigerator on its side so that the flies will not be trapped in the media. After 20 minutes or so check the cultures to make sure the flies are unconscious. Then tap the flies into a Petri dish. Using the camel’s hair brush, separate the flies into different groups (e.g., males and females; red eye and white eye, etc.) In preparing a new culture, five to ten pairs of flies should be used.

It is not difficult to distinguish between the male and female Drosophila. With a little practice, the difference can be seen without magnification (refer to Figure 1). The female is usually slightly larger than the male. The abdomen of the male appears to have a distinct black tip and a blunt posterior, while the female has a somewhat lighter and pointed posterior with bands visible almost to the tip. The male also has sex combs on the front legs and the female does not. As the female becomes older, her abdomen becomes distended with eggs, making identification even more simple.
Figure 1

To inoculate the culture jar with the flies, sweep the flies onto the end of a piece of filter paper. Lay the culture jar on its side and remove the plug. (Do not allow the flies to fall off the paper.) With the handle of the camel’s hair brush, push the end of the filter paper down into the medium (See Figure 2). Plug the jar. Allow the jar to remain on its side until the flies awaken. This will help to avoid the possibility of the unconscious Drosophila falling into any moisture and drowning.

Figure 2

It is good practice to check the characteristics of each strain under magnification when subculturing. There is always the possibility that a stray fly may enter the culture and cause loss of the strain’s purity.

Stocks should be subcultured every 2 – 4 weeks depending upon the temperature. Old cultures should be discarded after the new cultures become established in order to avoid contamination. At least two cultures of each strain should be kept in case one should be unsuccessful.

It is important to make sure that each culture jar is labeled with the type of fly and the date the jar was inoculated with the flies. The best way to do this is to use the shorthand notation for the particular strain of fruit fly in the jar. An example of a few shorthand notations would be: + = wild type (red eye), vg = vestigial wing, b p c = black purple curved. Small letters refer to recessive genes, capital letters refer to dominant genes (e.g., B = Bar eye). Letters separated by spaces refer to genes on the same chromosome (e.g., wem = eosin miniature). Letters separated by a semicolon (e.g., w;se – white sepia) refer to genes located on different chromosomes.

Therefore, a culture jar with a heldout strain of D. melanogaster on February 14 would have the following label:
2/14 Label

There are so many other notations and additional symbols that it would be impossible to list them here. All of WARD’S cultures are shipped with a label giving the name of the strain, the shorthand notation, and the chromosomes on which the genes occur.

Cultures of Drosophila should be kept at a relatively constant temperature closely approximating room temperature, no lower than 20°C (68°F) and no higher than 25°C (77°F). They should not be exposed to direct sunlight and may be kept entirely in the dark.

Embryonic development (following fertilization) takes place within the egg membrane. The egg hatches and produces a larva which feeds by burrowing through the medium. As the larva grows, it undergoes two molts so that the larval period consists of three stages (instars), the first instars being the newly hatched larva. The final larval stage or third instar may attain a length of 4.5 mm. The third instar stage, toward the end of the larval period, will crawl up the sides of the culture jar, attach itself to a dry surface (the jar, the filter paper, etc.) and form the pupa. After a period of time the adult or imago will emerge.

The duration of the above stages will vary with the temperature; at 20°C (68°F) the average length of the egg-larval period is eight days, while at 25°C (77°F) it is reduced to five days. Thus at 25°C (77°F) the life cycle may be completed in about ten days, while 20°C (68°F), fifteen days are required.

The sperm received by a female fly during mating is retained, serving to fertilize a number of eggs. Therefore, in an experimental cross between two different strains, virgin females must be used. This can be done quite simply by taking advantage of the fact that females do not mate before twelve hours from the time of emergence. If all the adult flies are removed from a culture with many pupae, all of the females collected within the next twelve hours will probably be virgin.

Experimental Crosses

Perhaps the best way to explain the technique used in making experimental crosses would be to trace the step-bystep procedure, using an example. Careful records should be kept of all experiments, giving dates of crosses, results of crosses, comments about technique, and results of the experiments.

For our experiment, we will use the wild type (+) strain and the vestigial wing (vg) strain.

  1. Select virgin vestigial (vg) females and place six in each of three culture bottles. Into each bottle place six wild type (+) males. Mark the date and the type of mating on each bottle.
    1_exp-cross.gif
  2. After seven days remove the parent flies from the mating bottle and discard. There should be many larvae in the bottle.
  3. When the flies begin to emerge, examine them and record the characteristics. This is the F1 generation. In this case, since the (vg) is recessive, all of the flies should exhibit (+) characteristics. (Note: If any (vg) flies appear in a bottle, one of the parent females was NOT a virgin and the culture should not be used in the rest of the experiment.)
  4. Place six males and six females of the F1 generation in each of three culture bottles. Mark each bottle with the date, the type of original mating, and the generation.
    4_exp-cross.gif
  5. Remove the F1 flies from the culture bottles after seven days and discard them.
  6. When the offspring of this cross (the F2 generation) begin to emerge, they should be killed (by over-anesthetizing), removed, and the sex and characteristics determined and recorded. Counting should go on until all flies have emerged and have been counted (usually about eight days, depending on the temperature). Counts should be made daily.
  7. The results should look like this:
    7_exp-cross.gif

The wild type flies outnumber the vestigial wing flies by about three to one. The sex of the flies does not appear to affect the ratio (in this case). The crosses can be diagrammed to predict this in a very simple manner.

Since the F1 generation is the result of two homozygous strains of flies, the wild type fly has only one possible characteristic in the gametes, wild (+). The vestigial fly also has only one possible type of gamete, vestigial (vg). The F1 generation can therefore be diagrammed:
F1 Diagram
The zygotes (fertilized eggs) all have both the wild and the vestigial genes. Since the wild type is the dominant factor, all of the F1 flies will appear to be of the wild type, even though they are not homozygous strains.

The F1 has TWO possible gametes, wild (+) or vestigial (vg). Therefore, the F2 would look like this in a diagram:
F2 Diagram
For every F2 fly with the vestigial characteristic, there will be three flies with the wild characteristic (one homozygous for wild and two heterozygous for wild).

As an additional experiment, you could try crossing F1 generation virgin females with pure wild or vestigial strains and try to predict the results.

A brief survey of methods for working with Drosophila is all that has been possible within the scope of this leaflet. The reader is urged to take advantage of some of the fine works available which deal with Drosophila and genetics in general for more detailed information.

This guide is also available in PDF format on wardsci.com.

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