Neurobiology, Biol 635: LABS

C161 fly from Williams & Smith 2002; confocal image by Richard Vogt Lab 1: Neural Anatomy of Drosophila
Fluorescent and Confocal Microscopy of Transgenic Fly Larvae (introduction to microscopy methods).

Part 1: Visualizing Neurons: GFP (Green Fluorescent Protein).

(NOTE: We will do this during the first week.)

Students will observe Drosophila larvae behaving, and examine the larval neural anatomy using fluorescent and confocal microscopy. Subsets of neurons in the nervous system of these flies express Green Fluorescent Protein (GFP). These flies were the result of enhancer trap experiments. A P-element (transposable element) carrying a reporter gene was introduced in the flies and allowed to randomly insert into the host chromosomes. In the flies of interest, nearby regulatory regions (presumably) induce GFP expression in tissue specific manner. The flies we will examine express GFP in the brain and motor neurons (D42) or in peripheral sensory neurons (C161). Student introduction to the microscopy techniques will be brief, but hopefully encouraging for students to choose an anatomically oriented project that will make use of this instrumentation.

Two papers describing use of C161 line:

Shepherd D, Smith SA (1996) Central projections of persistent larval sensory neurons prefigure adult sensory pathways in the CNS of Drosophila. Development 122, 2375- 2384.

Williams DW, Shepherd D (2002) Persistent larval sensory neurones are required for the normal development of the adult sensory afferent projections in Drosophila. Development 129, 617-624.

Part 2: Visualizing Neuronal Lineages.

(NOTE: This experiment requires an extended period of time ? students embarking on this project need to be aware of the importance of scheduling.)

The development of a complex animal from a single cell embryo involves many small sequential decisions. Asymmetries are established, cells become different and these different cells divide, establishing cell lineages which take on different roles. The nervous system derives from neuroblasts which in turn differentiate from ectodermal cells early in development. These neuroblasts divide, and their progeny establish the many neuronal lineages (neurons related by which neuroblasts they derive from). Members of different neuronal lineages play different roles within the nervous system.

Studies of cell lineages has often involved finding a way to visibly mark a specific cell; when that cell divides it passes its visible marking on to its progeny cells, and they pass the mark onto their progeny, and so on. For example, early developmental biologists would inject a colored die into a blastomere (one cell of perhaps an 8 cell embryo); by following the fate of the colored die (which cells ended up with it), it was possible to identify which parts of the body derived from that specific blastomere.

MARCM is a genetic method to label neuronal cell lineages in Drosophila. MARCM stands for "Mosaic Analysis with a Repressible Cell Marker", and was described in a paper by Lee and Luo in 1999:

Lee T, Luo L (1999) Mosaic analysis with a repressible cell marker for studies of gene function in neuronal morphogenesis. Neuron 22, 451-461.

Two papers below use the MARCM technique to study the sensory neurons in C161 flies (discussed above) and in neurons of the developing adult Drosophila during metamorphosis. The second paper used the specific reagents (genetic constructs) you will use in this exercise.

A MARCM experiment is very easy to do, and produces clusters of neurons which fluoresce green light under a fluorescent microscope... very cool.

Procedurally, you...

. combine males and females of two genotypes in a vial and let them mate and lay eggs for 2 hours;.
. remove the adults and let the eggs develop for 3 more hours at room temperature;
. place the vial in a 37^oC water bath for one hour;
. place the vial on the shelf and wait several days for development to progress;
. look at the flies (dissected) using a fluorescent microscope --- take pictures.

Technically, the way the experiment actually works is remarkably sophisticated, complicated, and probably pretty difficult to understand. I will try and explain this in steps, but you should also read the following short essay...

Luo L, Lee T, Nardine T, Null B, Reuter R (1999) Using the MARCM system to positively mark mosaic clones in Drosophila. Drosophila Information Service 82, 102-105.
1. A fly is constructed with the following genotype: (promotor)Gal4; UAS-GFP. In this fly, the promoter drives the expression of a transcription factor called Gal4, and Gal4 binds to and activates a regulatory site referred to as "UAS" (upstream activating sequence). Activation of the UAS site drives expression of GFP (green fluorescent protein) which fluoresces green when stimulated by blue light.
2. This fly also contains a gene encoding and expressing a protein called "Gal80"; Gal80 suppresses the action of Gal4. If Gal80 is expressed, no GFP is made and no green fluorescence can occur.
3. This fly also contains a complex of genes referred to as the FLP/FRT system; FLP is a transcription factor that activates the FRT site, which is situated adjacent to the Gal80 site. Further more, at least in our case, the FLP is driven by a "heat shock" promoter (hs). All this means is that when you raise the temperature of the animal to 37^oC, this activates the hs promoter which activates the expression of FLP which activates the FRT site.
4. So we make a bunch of fly embryos that have all this stuff in them. Procedurally this is really easy, since the genes have already been put in the flies, and all we have to do is take virgin females of one stain (FRT-Gal80) and mate them to males of another strain (FRT-UAS-GFP) and... POW... we have fly embryos that have all this stuff in them.
5. All the cells in the embryos we now have are capable of expressing GFP except for the one problem... all the cells are expressing Gal80 which is blocking the expression of GFP. We need to turn off Gal80 expression. We do this by activating the FLP/FRT system.
6. Normally, a cell has two copies of each chromosome called chromatids. In our case, the chromatids are different, one containing by FRT-Gal80 and the other containing FRT-UAS-GFP. This cell can not express GFP because Gal80 is present. During mitosis, the chromatids are duplicated and sort to produce two identical chromatid pairs, both pairs consisting of a FRT-Gal80 chromatid and a FRT-UAS-GFP chromatid. Like their mother, neither daughter cell would be able to express GFP, again because Gal80 is present. HOWEVER, AND HERE IS THE TRICK... if the FRT is activated during mitosis, it induces a recombination event (recombination normally only occurs during meiosis), creating one chromatid pair that contains only UAS-GFP and another chromatid pair that contains only Gal80. One of the resulting daughter cells now contains no Gal80, and suddenly is able to express GFP and fluoresce green light. And any additional cells produced by this daughter will also express GFP. Cool!!!

We are providing you with two different FLP/FRT systems to explore...

System 1 has the following genotypes:

strain 1:

strain 2:

crossing a virgin females of strain 1 with males of strain 2 will result in embryos with the following genotype:

hsFLP ; 42B-FRT, tubP-Gal80 / 42B-FRT, UAS-mCD8::GFP ; actin-Gal4/wt

System 2 has the following genotypes:

strain 1:

strain 2:

crossing a virgin females of strain 1 with males of strain 2 will result in embryos with the following genotype:

hsFLP, hsFLP ; 42B-FRT, tubP-Gal80 / 42B-FRT, UAS-mCD8::GFP ; wt/wt

Selecting virgin females.
You will need to select virgin females. This is easy to do, but must be done several times during the day, and you must be able to distinguish male and female adult flies. The life stages of a fly include larva (4 days, 3 instars); pupa (4 days) and adult (4 weeks). You need to select virgin females immediately after they emerge as adults from the pupal stage. This is quite easy, as an adult fly looks quite different than a pupa (wings, legs, etc ? looks like a fly). The trick is getting the females out before they have mated (lots of pesky males are in the same vial). This also turns out to be quite easy since the females will not mate until about 8 hours after they emerge (although I would not trust hours 7 and 8). And it is even easier since most of the flies emerge in the morning, after the lights go on.

Collecting virgin females in primarily an effort of knowing how old the females are, and this is done by frequently removing all adult flies from the vial. Ideally, you should remove all adults from the vial immediately after the lights go on. Then simply check back 6 hours later, collect all adults, separate males from females, put females in a vial and leave them for a couple of days to ensure no embryos are laid (an indication that you missed one). After confirming these are all virgins, they can be used for your MARCM experiment. You can not have too many virgin females.

Summary of experiment.

Collect virgin females.
Combine a large number of virgin females (strain 1) and males (strain 2) in a vial.
In the morning, transfer the flies to a new vial containing yeast granules. Allow these to lay embryos for 2 hours (check that embryos are on the surface of the food). Transfer all adults to a new vial (also containing yeast granules).
Allow embryos to develop 3 hours.
Apply heat shock (place vial in 37^oC water bath for 1 hr).
Place vial in fly rack to allow development to continue.
Examine fly larvae at 3rd instar (~3 days later), or pupae and adults at appropriately later stages)

We will show you how to dissect larvae and how to use the fluorescent and/or confocal microscopes.

What to look for...
Initially look for GFP fluorescent neuronal clusters in 3rd instar larvae or pupal CNS. Photograph these using the fluorescent microscope and make 3-dimensional images using the confocal microscope. Document as many different neuronal architectures (different looking neurons) as you can.

More sophisticated projects can involve studies of the role of hormones in regulating neuronal development, or the labeling of neurons in imaginal discs (presumptive adult structures). But first things first ? learn how to do the experiment and visualize specific neuronal lineages!

from www.proweb.org/kinesin/ WTKHClarvae.html Lab 2: Associative Learning by Drosophila larvae
An introduction to neuroplasticity and data analysis.

Plasticity. Organisms must be able to adjust to changing conditions. Nervous systems are designed to be flexible (plastic). When a nervous system makes a decision, it assesses external information via sensory inputs, and compares this with internal stored information (memories of previous experiences), and an understanding of the context of the situation. Based on all this (and perhaps colored by the factors such as how much sleep or alcohol the organism has recently consumed), an appropriate response is determined and implemented. This is pretty complicated, involving many mechanisms.

Drosophila larvae offer an opportunity to explore this operation. Larvae will crawl towards or away from certain odors (that may indicate the presence of food or of something to avoid), and will display a preference for staying in areas of food (if given a choice). We can manipulate these responses; the fly nervous system is capable of developing different interpretations of a specific stimulus.

Chemo-orientation. Adult flies can smell volatile odors with their antennae (with sensory neurons on their antennae) and will fly up an odor plume to the source of the odor. You could make a small wind tunnel using some tubing (say 4" diameter), putting an odor (banana) at one end and some adult flies at the other. If you create a bit of air movement that blows the banana smell towards the flies, they should fly up the tube to the banana. You could create the air movement using a fan, or by setting the fly-end of the tube in a fume hood which would suck the banana odor down the tube. You could also construct a behavioral assay that would allow you to quantify the attraction or repulsion of flies to specific odors, such as in these three videos:

1. Beetle Olfactometer (Tania Suerda, IIQAB, CSIC, Barcelona, 2/20/02) (1:42)
2. Drosophila Attractants & Repellants (Anna Lisa Balloi, ALNARP, 4/2002) (8:40)
3. Drosophila larval chemoreception assay (28:00)

Two companies developing equipment for monitoring behavior/physiology:

In these studies, we are not testing whether or not the animal will respond behaviorally to the odor. This is much more than just determining if the animal is smelling the odor. The animal might smell an odor (i.e. its chemosensory neurons might be stimulated), but it might not be interested in that odor and therefore may not respond. Or the animal might only be interested in responding to the odor stimulant in a specific context.

Drosophila larvae can also smell odors and taste chemicals, using chemosensory neurons on their head (review Guide to Larval Anatomy). In this exercise, we will mess this tiny animal's even more tiny brain. We will create a context where the animal will respond to an odor in a way that is independent of the odor and, depending on how we do it, different than the "normal" respose. We will teach the animal to associate a smell with a reward (sugar) or punishment (salt), and then test if the animal will approach or avoid the odor in a manner that is independent of the odor itself. This is called "associative learning". We are going to teach a fruit fly maggot a new trick! (Food for thought?)

The three papers below will be a basis for these experiments. The first paper describes the experimental set up, and the second and third papers describe experiments done with this assay.

PDF Scherer S, Stocker RF, Gerber B (2003) Olfactory learning in individually assayed Drosophila larvae. Learning and Memory 10, 217-225.
[Assay details; includes explanation of odor choice.]
PDF Gerber B, Scherer S, Neuser K, Michels B, Hendel T, Stocker RF, Heisenberg M. (2004) Visual learning in individually assayed Drosophila larvae. J. Exp. Biol. 207, 179-188.
[Assay details.]
PDF Neuser K, Husse J, Stock P, Gerber B (2005) Appetitive olfactory learning in Drosophila larvae: effects of repetition, reward strength, age, gender, assay type and memory span. Animal Behavior, 69, 891-898.
[Details / methodology of associative learning assay (petri dish).]
PDF Michels B, Diegelmann S, Tanimoto H, Schwenkert I, Buchner E, Gerber B (2005) A role for Synapsin in associative learning: The Drosophila larva as a study case. Learning and Memory 12, 224-231.
[Use of associative learning assay (petri dish), and as well showing surprisingly minimal effects of synapsin deletion on learning. ]
PDF Hendel T, Michels B, Neuser K, Schipanski A, Haum K, Sokolowski MB, Marohn F, Michel R, Heisenberg M, Gerber B J. (2005) The carrot, not the stick: appetitive rather than aversive gustatory stimuli support associative olfactory learning in individually assayed Drosophila larvae. Comp. Physiol. A 191, 265-279.
[Study of fructose, salt and quinine on associative and non-associative learning of Drosophila larvae, petri dish assay.]

We will read the papers together in class and discuss the experiments, and then we will perform and extend these studies. One experiment I am especially interested in is testing whether or not the animals can distinguish odor molecules that have been classified to belong to the same class, per the following paper:

PDF Stensmyr MC, Giordano E, Balloi A, Angioy A-M, Hansson BS (2003) Novel natural ligands for Drosophila olfactory receptor neurones. Journal of Experimental Biology 206, 715-724.

You can read more about insect chemodetection from the following web page: http://www.biol.sc.edu/~vogt/pdf/olf/olf-read-list.html.

Lab 3: Cricket Circal System
Sensory Integration (electrophysiology and behavior).

Papers describing the cricket circal system:

The cerci and abdominal giant fibres of the house cricket, Acheta domesticus. I. Anatomy and physiology of normal adults. Edwards JS, Palka J (1974) Proceedings of the Royal Society of London, Series B, Biological Sciences. 185, 83-103.
Receptive fields of cricket giant interneurons are related to their dendritic structure. Bacon JP, Murphey RK (1984) Journal of Physiology 352, 601-623.

Sensory systems gather environmental information, and present this information to the central nervous system in a manner that the CNS can interpret. Part of this presentation is the electrical activity of the sensory neurons. But a large part is also the reconstruction of a spatial interface that in some manner reconstructs the spatial qualities of the environment, as they relate to the sensory modality in question. The Cricket Circal System is an elegant example of the organization of such an interface. Crickets use their cerci to determine the direction from which wind is blowing. Many sensory hairs line the cerci and are capable of being moved in restricted directions: some move from wind coming from behind, while others move from wind from the right, from the left or from the front. The source of the wind might be the pressure wave riding the front of a frog's tongue, whipping out to capture the cricket. Sensory neurons from the sensory hairs on the cerci project into the terminal abdominal ganglion (the circal ganglion), where they make synaptic connections with "Giant Interneruons". The giant interneurons have cell bodies located within the circal ganglion: their dendrites fill specific regions within the neuropile of the ganglion, and their axons project up the nerve cord (anterior) to the thoracic ganglion where they synapse on neurons which control the jumping legs. When the frog tongue comes near, the cricket's circal system conveys both its immanent arrival and the direction from which it is coming, communicates this information very quickly to the jumping system, and the cricket jumps out of the way... maybe...

This lab revolves around this cricket circal system. You will mount and dissect a cricket, place hook electrodes under its abdominal ventral nerve cord, and perform experiments which reveal aspects of sensory integration.

The accompanying videos present a scientific paper that describes this system, and the methodology of how to prepare the crickets for the dissection.

You should maintain a detailed record of your efforts, write a brief report and present your effort to the class on the second Thursday. This presentation can be in powerpoint, a web site, video, or a combination.

Lab 4: Cricket Visual System
Sensory Physiology (electrophysiology and behavior).
(image from: http://www.technovelgy.com/ct/Science-Fiction-News.asp?NewsNum=1638)

Teaching insect retinal physiology with newly designed, inexpensive micromanipulators. (2006) Krans J, Gilbert, C, Hoy R Adv. Physiol. Educ 30, 254-261.

5th instar Manduca, photo by Richard Vogt Lab 5: Manduca Heart
Modulation of Electrically Excitable Tissue by Neuropeptides and Biogenic Amines (electrophysiology and neuropeptides).

Aids:

Nerves and muscles share the property of electrical excitability, though differ in their embryonic origins (nervous tissue derives from ectoderm while muscle derives from mesoderm). Insects have both skeletal muscle and smooth muscle. Insect skeletal muscle is striated, like ours, but is not capable of generating self propagating action potentials (unlike ours); insect skeletal muscle has voltage sensitive K⁺ channels, but lacks voltage sensitive Na⁺ channels. Insect smooth muscle, however, is capable of generating self propagating action potentials. An excellent example of such a muscle is the dorsal heart or dorsal vessel. In this laboratory exercise you will electrically record peristaltic contractions of the larval heart of the moth Manduca sexta and study its modulation by Cardioacceleratory Peptides (CAPs, neuropeptides) and serotonin and octopamine (biogenic amines).

While insects have open circulatory systems, they have many pumps or pulsatile organs to move their blood (hemolymph), as well as structural components that help direct the flow of blood (see this paper by Gunther Pass reviewing these accessory hearts). Insect hemolymph transports nutrients, cellular waste and hormonal signals; gas exchange occurs via the separate tracheal system.

Quoting from Chapmann, (1998, p. 94) ... "The dorsal vessel runs along the dorsal midline, just below the terga, for almost the whole length of the body although in the thorax of adult Lepidoptera ... it loops down between the longitudinal flight muscles... It may be bound to the dorsal wall or suspended from it by elastic filaments. Anteriorly it leaves the dorsal wall and is more closely associated with the alimentary canal, passing under the brain just above the esophagus. It is open anteriorly, ending abruptly in most insects... Posteriorly, it is closed. The dorsal vessel is divided into two regions: a posterior heart in which the wall of the vessel is perforated by incurrent and sometimes also by excurrent openings (ostia), and an anterior aorta which is a simple, unperforated tube. The heart is often restricted to the abdomen... The wall of the dorsal vessel is contractile and usually consists of one or two layers of muscle cells with a circular or spiral arrangement. Longitudinal muscle strands. ... The incurrent ostia are vertical, slit-like openings in the lateral wall of the heart. ...many Lepidoptera have seven or eight. ... In the silkworm Bombyx, only the hind lip of each ostium is extended as a flap within the heart." In most insect species ... the heart is innervated by nerves running round the body wall from the segmental ganglia. In ... larval Lepidoptera ... branches of the segmental nerves combine to form a lateral cardiac nerve running along each side of the heart."

The heart of M. sexta is both myogenic and neurogenic. The heart can and does contract on its own in the absence of neuronal input (myogenic), but the heart also receives neural input (see Wasserthal paper below). The heart's contractile activity is modulated by the biogenic amines octopamine (neural?) and serotonin (octopamine mimic?) and by the neurohormone CAP (cardioacceleratory peptide). These substances influence the heart in the same direction, increasing its rate of contraction. However octopamine is thought to act via a cAMP pathway while CAP is thought to act via an IP3 pathway.

First Effort Experiment (Everyone should do this as soon as possible):

2. Still quickly, pin the animal to the Petri dish (sylgard filled). Use two forceps to stretch apart the epidermis at the anterior end first, and affix two pins. The work backwards, stretch and pin, stretch and pin, etc. 3. Look at your preparation. Rinse with saline. Notice and identify as many bits and parts as possible.

4. Carefully but firmly, grab the intestine at the anterior end with forceps and cut on the anterior side of the forceps. Gently lift the gut up while tearing away tracheae using a glass probe. Finally, cut the gut free and discard (gut enzymes may destroy your preparation).

5. Look at your preparation. Rinse with saline. Notice and identify as many bits and parts as possible. Notice the ventral nerve cord with its ganglia. Notice the heart as a clear region running along the "dorsal midline". This may be difficult to see in older animals if fat body is excessive.

6. Set saline to drip slowly onto animal (dishes are at an angle; drip onto upper end; have drip-tip as close to animal as possible.

7. Find a region of heart that is "beating" and gently place the tips of the silver wire electrodes against this region. Place the ground (discharge) electrode somewhere in contact with the tissue.

8. Record slow contractions.
Amplifier settings should be: Gain 10k, Low pass filter = 1, High pass filter = 1k
Computer settings might be: samples per second = 100; Y-axis = +/- 1 volts; Samples per sweep = 500; X-axis = 500 (will record 5 seconds of data @ 100 samples per second).

9. Try recording with longer recording times (same samples per second but increase samples per sweep and X-axis range).

10. EXPERIMENT:

Manduca

You will drip the homogenate onto the heart, comparing contraction frequencies (and amplitudes) before and after treatment, as well as following rinsing with normal Manduca saline.

Do the following. Set to record for 2 minutes. Flush with saline for 1 minutes, then turn off saline. Start recording. At 30 seconds, pipette 200 ul Manduca saline onto uphill end. After recording has stopped (and been saved), flush with saline for 1 minute. Stop saline, start recording. At 30 seconds, pipette 200 ul of nerve cord homogenate at uphill end. After recording has stopped (and been saved), flush with saline for 1 minute. Stop saline, start recording and at 30 seconds pipette 200 ul of Manduca saline onto uphill end. After recording has stopped (and been saved), compare the three recordings. Calculate the Frequency of Heart Rate under each of the three conditions.

After performing the "First Effort" excersize below, further investigation with this prepration affords the opportunities to explore several aspects, including:

References:

General References:

Chapman RF (1998) The Insects, Structure and Function, 4th ed. Cambridge University Press, Cambridge.
Hoyle, G (1983) Muscles and Their Neural Control. John Wiley & Sons, New York.

Pulsatile Organs in Insects ? Evolution of Insect Hearts:

Pass G (2000) Accessory pulsatile organs: evolutionary innovations in insects. Annual Review of Entomology 45, 495-518. PDF

Anatomy and regulation of heart:

Heinrich B (1970) Nervous control of the heart during thoracic temperature regulation in a Sphinx moth. Science 169, 606-607. PDF
Heinrich B (1971) Temperature regulation of the Sphinx moth, Manduca sexta. I. Flight energetics and body temperature during free and tethered flight. J. Exp. Biol. 54, 141-152. PDF
Heinrich B (1971) Temperature regulation of the Sphinx moth, Manduca sexta. II. Regulation of heat loss by control of blood circulation. J. Exp. Biol. 54, 153-166. PDF
Sanger JW, McCann FV (1968a) Ultrastructure of the myocardium of the moth, Hyalophora cecropia. J. Insect Physiol. 14, 1105-1111.
Sanger JW, McCann FV (1968b) Ultrastructure of the moth alary muscles and their attachment to the heart wall. J. Insect Physiol. 14, 1539-1544.
Wasserthal LT, Wasserthal W (1977) Innervation of heart and alary muscles in Sphinx ligustri. Cell Tissue Res. 184, 467-486. PDF

Cardioactive Peptides:

Platt N, Reynolds SE (1985) Cardioactive peptides from the CNS of a caterpillar, the tobacco hornworm, Manduca sexta. J. Exp. Biol. 114, 397-414. PDF
Tublitz NJ, Truman JW (1985) Intracellular stimulation of an identified neuron evokes cardioacceleratory peptide release. Science 28, 1013-1015. PDF
Tublitz NJ, Truman JW (1985a) Insect cardioactive peptides. I. Distribution and molecular characteristics of two cardioacceleratory peptides in the tobacco hawkmoth, Manduca sexta. J. Exp Biol. 114, 365-379. PDF
Tublitz NJ, Truman JW (1985b) Insect cardioactive peptides. II. Neurohormonal control of heart activity by two cardioacceleratory peptides in the tobacco hawkmoth, Manduca sexta. J. Exp Biol. 114, 381-395. PDF
Tublitz NJ (1989) Insect cardioactive peptides: neurohormonal regulation of cardiac activity by two cardioacceleratory peptides during flight in the tobacco hawkmoth. J. Exp Biol. 142, 31-48. PDF
Tublitz NJ, Broadie KS, Loi PK, Sylwester AW (1991) From behavior to molecules: an integrated approach to the study of neuropeptides. Trends Neurosci. 14, 254-259.
Tublitz NJ, Allen AT, Cheung C, Edwards KK, Sylwester AW, Reynolds SE (1992) Insect cardioactive peptides in Manduca sexta: a comparison of the biochemical and molecular characteristics of cardioactive peptides in larvae and adults. J. Exp. Biol. 165, 265-272. PDF
Prier KR, Beckman OH, Tublitz NJ (1994) Modulating a modulator: biogenic amines at subthreshold levels potentiate peptide-mediated cardioexcitation of the heart of the tobacco hawkmoth Manduca sexta. J. Exp. Biol. 197, 377-391. PDF
Smits AW, Burggren WW, Oliveras D (2000) Developmental changes in in vivo cardiac performance in the moth Manduca sexta. J. Exp. Biol. 203, 369-378. PDF

Manduca Saline (from: Tublitz & Truman, 1985, J. Exp. Biol. 114, 365-379; 381-395)

This will probably precipitate if you make it at once. I suggest makingit as follows.

Make up 200 ml at a time working solution, in the following order, while stiring or mixing):
20 ml solution #1
120 ml H2O
20 ml solution #2 (NaPO4)
20 ml solution #3 (Na2PO4)
20 ml solution #4 (CaCl2 - add very slowly while stiring).

photo by Richard Vogt Lab 6: Crayfish
Electrophysiology of Sensory and Motor Activities (electropyhysiology and behavior).

LAB OPTIONS:
1. Swimmeret Rhythms and Synaptic Transmission.
2. An Investigation of Abdominal Stretch Receptors.
3. Innervation of Opener and Closer Muscles in Crayfish Claw: Cybernetic Crayfish
4. Crayfish labs @ Smith College (by Richard Olivo)

Fresh water crayfish have been a model organism for neurobiological studies since the late 1800s. I have provided you with details for three projects, investigating (1) sensory neurons (muscle stretch receptors), (2) synaptic innervation of abdominal muscles and recording of swimmeret activity in motor neurons, and (3) neuronal control of opener and closer muscles in the crayfish claw. Before you consider any of the projects, I urge you to spend some time looking at and considering these animals before proceeding.

A very excellent general web resource for these exercises are Dr. Robin Cooper's pages (University of Kentucky), especially:
General Projects and Sensory Projects .

Animals are purchased from Carolina Biological (telephone order).

Crayfish top
1. Swimmeret Rhythms and Synaptic Transmission

WHY?

Brian Malloney

Using the associated laboratory description, you can also investigate certain aspects of neuromuscular synaptic transmission.

Resources:

Laboratory #1 Guide including recipe of Crayfish Saline, description of suction electrode construction, and description of dissection suitable to exercises 1 and 2 (stretch receptor).
IMAGES (PowerPoint): Showing dissected crayfish, electrodes and ganglia, and recording dishes.

Crayfish top

2. An investigation of abdominal stretch receptors.

In the crayfish, there are stretch receptors located in small muscles which span segments in the dorsal (upper) part of the abdomen. The nerves containing the stretch receptor axons run down the sides of the animal and travel to the ventral nerve cord.

Do the following...

Option 1.

B. Use your suction electrode to sample activities of nerves while you raise the tail (lifting on telsons).

Option 2

B. Peel away the ventral cuticle (along with the nerve cord), and pull out the large muscle mass. DO NOT damage the tail (telson) or any of the musculature / tissue attached to the body wall.

C. Now you have a seemingly empty shell which in fact still contains many muscles. Immerse this tail under Crayfish Ringers (physiological saline). Look carefully along the cut edge of the cuticle (body wall). You can see very small nerve endings (cut by your scissors), one for each segment, on each side of the body.

D. Use the suction electrode and draw a bit of nerve into the tip of the electrode. Lifting the tail should induce volleys of action potentials in these nerves.

Resources:

Laboratory borrowed from Biology 305 (Comparative Animal Physiology), instructor Reagan Lake (University of Maryland, Baltimore County).
- Lab description.
- Lab PowerPoint
  .
PhD Dissertation of Edwin Furshpan (1955, California Institute of Technology): Studies on certain sensory and motor systems of decapod crustaceans.

Crayfish top
3. Innervation of Opener and Closer Muscles in Crayfish Claw: Cybernetic Crayfish

Resources:

PAPER: The triple innervation of crayfish muscle and its function in contraction and inhibition. van Harreveld A, Wiersma CAG (1937) Journal of Experimental Biology, 14, 448-461.
TERM PAPER: The Crayfish: background and related neurological research pertaining to Cheliped (Claw) research. by student Todd Stock, Neuro 635, 5/97
LAB REPORT: Activation of opposing muscle groups in the crayfish claw. by Todd Stock, Neuro 635, 5/5/97
IMAGES (PowerPoint): Showing mounted claw, re. experiments by student Tod Stock, 5/97 (Neuro 635)

Neurobiology LABS Flys, Crickets, Moths & Crayfish

Part 1: Visualizing Neurons: GFP (Green Fluorescent Protein).

Part 2: Visualizing Neuronal Lineages.

Neurobiology LABS

Flys, Crickets, Moths & Crayfish