Informative title for protocol

Kylie Lennon, Andrew D. Nguyen

YYYY-MM-DD format

Objectives and Questions

Comparing diapause duration with circadian rhythum.

natural history

Origin

Rhagoletis pomonella (Endopterygota:Tephritidae), commonly known as the apply maggot fly is native to North America. Primitively, they ate hawthorn (Crataegus) berries. When apples were introduced to North America in the 17th century, Rhagoletis began to use apples as a host. The large size of an apple reduces the ability of parasitoid wasps penetrating the maggots. However, apples ripen earlier in the season, beginning in early summer.

Physical Description

Emergence Habits

Beginning in early summer, adult Rhagoletis begin emerging. After about eight to ten days, females use their ovipositor to inject her eggs into the fruit. Once the eggs hatch, the maggots remain in the fruit. They continue to develop until the fruit falls from the tree.

Rhagoletis Project Workflow

Field Collections

Locations

dates

Collecting larvae and pupae

Reagents

Protocol

Prepare tubes for holding fruit with maggots

  1. Put the metal mesh into tubs, and pour the fruits onto the metal mesh
    • 1-2 layers of fruit
    • Use paper towels to clean up juices at the bottom of the tubes as the fruit begins to rot.
  2. Label tubes by site and host of fruit.
  3. Once larvae/pupae begin to exit the fruit, remove the metal mesh from the tub.
  4. Pick up the larvae/pupae gently with your fingers or forcepts, and place in a petri dish
    • less than 100 per petri dish
  5. Place the fruit and metal mesh back into the tubes.
  6. Split the larvae/pupae into several other petri dishes containing wet vermiculite.
    • less than 100 per petri dish
    • To prepare vermiculite, add water until all of it is wet. Water should not be dripping from dish.
  7. Label petri dish with: site, race, collection date of larvae/pupae, fridge date, number specimens per dish.
    • Fridge date is 13 days after collection date.
  8. Seal each petri dish with tape, and place dishes into a container with saturated potassium chloride (KCl) solution to stimulate humidity (85%).
    • KCl Solution: 1L water with 350g of KCl

Weighing and Purging

On days 10 and 14, the mass of each pupae will be taken. Then, each syringe will be purged. During this process, air is circulated through drierite and water with a pH of 4.0, which removes the carbon dioxide present. This allows the amount of carbon dioxide produced by each pupae to be measured approximately 24 hours later.

Weighing

Reagents

Protocol

  1. 10 days after the collection date, remove pupae from the vermiculite. Place in a clean petri dish.
    • Used dampened paper towel to clean all vermicute off of the pupae.
  2. Weigh pupae using an analytical balance to the nearest 0.001 mg.
  3. Place each pupae into separate air tight syringes.
    • Each syringe should have a colored piece of tape and number.
  4. Twist valve onto syringe.
    • Make sure the knob denoted with (off) is facing away from the syringe. This ensures the pupae will not fall out.
  5. Record the number, tape color, site, and mass for each pupae. Note: Four blanks should be included in each cohort.

Image: Example of syringe

Purging

Setup
A pump is attached to tubes which run through drierite and into an erlenmeyer flask containing H20 with a pH of 4.0. There are two valves that create a continuous flow of air through the setup. Another tube in the flask is used to inject clean air into the syringes containing the pupae.

Image: Purging setup

Protocol

  1. Plug in pump.

  2. Turn the two valves 90 degrees so the closed knob, denoted with (off), is in the downward position.
    • This will allow continuous clean air flow through the drierite and acidic water.
    • Make sure pupae is in a location where it will not be smashed when valve is rotated.
  3. Screw the valve to the end of the suction tube.
  4. Rotate the valve 90 degrees counterclockwise. Wait a few seconds for the air to be cleaned. Then, rotate an additional 180 degrees counterclockwise.
  5. Inject 1ml of air into the syringe.
  6. Rotate valve 90 degrees clockwise and unscrew from tube.
  • This process should be repeated for all of the pupae in the cohort. Record the first and last purging times.

Respirometry

Background

Respirometry measures the metabolic rate of an organism by measuring CO2 production.

Reagents

Setup

ExpeData software is installed into a computer. Tubes run through the setup, which allows for complete air circulation. A pump is connected to drierite and aspirite columns. These chemicals absorb the CO2 in the air. Then, air is circulated through a tube of magnesium to remove any H2O present. Two mass flow controllers set the flow rate. Two licor machines (6262 and 7000) are used to inject the air. (* need to come back) Image: Licor 6262 (top) and Licor 7000 (bottom)

Protocol

  1. Turn on pump. Wait approximately five minutes to remove the residual CO2.
  2. Label which color the licors correspond too.
    • Ex: Red syringes are injected into Licor 6262 and orange into Licor 7000.
    • Use the same licor and color setup for respirometry day 15.
  3. Screw syringe into the valve connected to the proper licor.
  4. With the closed portion of the valve facing downward, twist the knob clockwise 270 degrees. *Off should be on the right-hand side. This will allow no air to escape from the syringe.
  5. Inject the 1 ml of air.
  6. Turn the knob clockwise 90 degrees, returning to its original position.
  7. Unscrew the syringe and place aside.
  8. Place a marker in between each sample.
    • Allow the CO2 curves to level out before placing marker.
  9. Repeat for the remaining syringes.
  • Steps 3-7 should be completed for both licors simaltaneously. Two syringes should be inserted, injected, and removed one after the other before inserting a marker. This will be more time efficient.

Treatments

Reagents

  • 96 Well Clear Flat Bottom Polystyrene not treated Microplates Costar#3370
  • Conical Screw Cap Microcentrifuge Tubes (2ml) #02-681-344
  • Microcentrifuge Tubes caps #02-681-368

Protocol

  1. Place each each pupae into its randomly assigned cohort treatment.
    • This should be done immediatly after day 15 respirometry is performed.
  • RT (room temperature)
    • Pupea is placed in assigned wells of microplates.
    • The microplates are kept in approximately 23 degree celcius conditions. They are placed in tubs containing KCL.
  • SO (stimulated overwinter)
    • Pupea is placed in assigned wells of microplates.
    • Microplates are kept in a tub in the fridge.
  • Each plate should be labeled with the day of treatment, type of treatment, collection date, cohort day, and cohort date. Image:A example of a labeled microplate
  • GC (genetic control)
    • Pupea is placed in microcentrifuge tube. Its unique individual id should be written on the centrifuge tube cap.
    • Placed in -79 degree Celcius freezer. Image: This is the setup of RT cohorts and SO cohorts once removed from the fridge. The tubs are organized by apple, hawthorn, skipped cohorts, and stimulated overwinter.

Eclosions

Background

  • Once the fly or parasitoid wasp has completed development, it will emerge from its pupae covering. At this point the adult needs to be transferred from the microplate into a tube containg food and water.

Reagents

  • Fly handling cage
    • Its a clear plastic box. One side is composed of mesh material, allowing for ventilation. The front side contains an open circle surrounded by netting.
  • 15 ml Polystyrene Conical Tube #352099
  • Food (Combination of brown sugar and yeast)

Protocol

Checking for Eclosions

  1. Check daily to see if an organism has emerged from a pupae shell.
  2. If there is an eclosion, make a table consisting of:
    • Host, cohort day, well ID, type of organism, trikinetics, and trikentics entry time.
    • the trikenetics and entry time will be recorded after transfer of organism.

Prepare the tubes with food and water

  1. Take a cotton ball and split it in half.
  2. Attach a tiny piece of paper (< cm in length) to the cotton using tape.
  3. Place a few drops of the food onto the paper.
    • Use enough food to fill paper without getting food onto the cotton.
  4. Insert the cotton with food into the open tube.
  5. Taking a conical tube, fill it up completely with water.
  6. Place a small piece of cotton at the top of the tube.
    • Make sure the cotton is wet but will not allow water to escape.
  7. Insert water tube into the other half of cotton ball. Place into other end of open tube.
  8. Using a piece of tape, label the tube with the individual id of the organism.

Tranfer of organism

  1. Place microplate and prepared tube into fly handling cage.
    • It’s beneficial to flip over the microplate top, and remove the cotton with the food.
  2. Using a bendable tube with cut pipette tips on each side, place mouth on one end of tube. Place the other end of tube over the organism and inhale.
    • Caution: Inhaling with too much force could lead to swallowing the organism. It is recommended to place a piece of mesh material around the mouth tip.
  3. Exhale, moving the organism into the prepared tube.
  4. Close the tube opening with the cotton, and remove from cage.

Trikinetics

Background

Trinkinetics systems measure the circadian rhythms and sleeping patterns of various types of small animals. The animal is placed in a tube where infared beam arrays continuously monitor the animal’s movement. In this experiment, once a fly or wasp ecloses, they are placed into the trinkentics system. They will be reared in 16 hours of light and 10 hours of dark increments. Some remain there until death. Others are taken out after eight days, and placed into a Free Run Trikentics system. The same type of monitor is used to measure the organisms activity level. However, the free runs are continuously kept in a dark room. The activity levels will be compared between the light-stimulated and free run trials.

Image: One of the trinketics monitors in the light-stimulated room.

Reagents

  • Locomotor Activity Monitor LAM25

Protocol

  1. Place the tube containing the organism into one of the trinkentics positions. Record the position number, Individual ID, and time of entrance.
    • If more than one monitor is being used, the monitor number should also be noted.
  2. Check daily for deaths. Change the food or water if necessary.
    • If organism dies, remove from the monitor. Place in microcentrifuge tube with the individual ID on the cap. Place into the gentic control freezer.
  3. Transfer specific individuals into new free run monitor positions.
    • This should be done at night.
    • The transferred individuals are prechosen based on their ecolosion date. (8 days)

References