Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica

Summary

In animals with large identified neurons (e.g. mollusks), analysis of motor pools is done using intracellular techniques^1,2,3,4. Recently, we developed a technique to extracellularly stimulate and record individual neurons in Aplysia californica⁵. We now describe a protocol for using this technique to uniquely identify and characterize motor neurons within a motor pool.

Abstract

In animals with large identified neurons (e.g. mollusks), analysis of motor pools is done using intracellular techniques^1,2,3,4. Recently, we developed a technique to extracellularly stimulate and record individual neurons in Aplysia californica⁵. We now describe a protocol for using this technique to uniquely identify and characterize motor neurons within a motor pool.

This extracellular technique has advantages. First, extracellular electrodes can stimulate and record neurons through the sheath⁵, so it does not need to be removed. Thus, neurons will be healthier in extracellular experiments than in intracellular ones. Second, if ganglia are rotated by appropriate pinning of the sheath, extracellular electrodes can access neurons on both sides of the ganglion, which makes it easier and more efficient to identify multiple neurons in the same preparation. Third, extracellular electrodes do not need to penetrate cells, and thus can be easily moved back and forth among neurons, causing less damage to them. This is especially useful when one tries to record multiple neurons during repeating motor patterns that may only persist for minutes. Fourth, extracellular electrodes are more flexible than intracellular ones during muscle movements. Intracellular electrodes may pull out and damage neurons during muscle contractions. In contrast, since extracellular electrodes are gently pressed onto the sheath above neurons, they usually stay above the same neuron during muscle contractions, and thus can be used in more intact preparations.

To uniquely identify motor neurons for a motor pool (in particular, the I1/I3 muscle in Aplysia) using extracellular electrodes, one can use features that do not require intracellular measurements as criteria: soma size and location, axonal projection, and muscle innervation^4,6,7. For the particular motor pool used to illustrate the technique, we recorded from buccal nerves 2 and 3 to measure axonal projections, and measured the contraction forces of the I1/I3 muscle to determine the pattern of muscle innervation for the individual motor neurons.

We demonstrate the complete process of first identifying motor neurons using muscle innervation, then characterizing their timing during motor patterns, creating a simplified diagnostic method for rapid identification. The simplified and more rapid diagnostic method is superior for more intact preparations, e.g. in the suspended buccal mass preparation⁸ or in vivo⁹. This process can also be applied in other motor pools^10,11,12 in Aplysia or in other animal systems^2,3,13,14.

Protocol

1. Preparation of Recording Dish

1. During the force transducer experiments, the buccal ganglia, cerebral ganglion, and buccal mass are placed in a round Pyrex dish that is specialized for force studies.
2. To induce ingestive-like patterns in the experiments, we need to apply the non-hydrolyzable cholinergic agonist carbachol to the cerebral ganglion¹⁵. To avoid direct contact from carbachol onto the buccal ganglia and buccal mass, separate chambers are needed to isolate the cerebral ganglion from the buccal ganglia and the buccal mass (Figure 1).
3. Since the buccal mass is much thicker than the buccal ganglia, they will not be placed on the same level. Therefore, this dish should have a back chamber for the cerebral ganglion (area A in Figure 1), a middle platform for the buccal ganglia (area C in Figure 1), and a much deeper front chamber for the buccal mass (area D in Figure 1).
4. To create this dish, begin with a round 100x15 Pyrex dish (15 mm high, 100 mm in diameter). Construction of the dish will require several pours of Sylgard. Follow the instructions provided with the Sylgard product. Sylgard must be allowed to set between different pours.
5. The first pour is to create the highest level of Sylgard in the dish (area B in Figure 1), which is the wall between the middle platform and back chamber.
6. Use two modeling clay backings to isolate the area for Sylgard wall (area B in Figure 1). Coat the modeling clay backings with plastic wrap where they will contact the Sylgard for easier removal. Ensure tight seals at the edges, where modeling clay will contact the dish, to minimize leakage.
7. Pour Sylgard into the portion between the two modeling clay backings nearly up to the top of the dish. Let the Sylgard fully set overnight. Keeping the dish in a warm place will induce faster setting. Remove the modeling clay backings and clean up any clay residue on the Sylgard.
8. Next, the back chamber (area A in Figure 1) and the middle platform (area C in Figure 1) should be poured.
9. Place a modeling clay backing about 5 mm away from the front surface of the Sylgard for the section of the middle platform (area C).
10. Pour Sylgard into the sections for the back chamber (area A) and middle platform (area C) up to a height of approximately 3-5 mm below the top level of the first Sylgard wall (area B). The Sylgard surface of the back chamber should be slightly lower than that of the middle platform to avoid leakage from the back chamber containing carbachol to the middle platform. Again, let the Sylgard fully set overnight and then remove the modeling clay backing.
11. The final step is to cut a notch in the middle of the Sylgard wall to provide a channel for the cerebral-buccal connectives (CBCs) to go through between the middle platform and back chamber. The width of this notch should be approximately 3-4 mm, which is wide enough for the CBCs. The bottom of the notch should not be lower than the Sylgard surface of the middle platform to prevent leakage. A scalpel blade can be used to cut the notch.

2. Electrode Preparation

1. Pull extracellular glass electrodes from single-barreled capillary glass using a Flaming-Brown micropipette puller as described by McManus et al.⁸ in section 3.1. With the FT345B filament in the puller, our typical program settings are Heat 480, Pull 50, Velocity 13, and Time 20, but note that settings will be different for different filaments. This program creates the electrodes in a single pull with no fire-polishing stage. The size of the electrode tip should be smaller than the size of the cell bodies. For the motor neurons ranging from 50 μm to 400 μm in soma diameter, the inner diameters of the extracellular glass electrodes should be about 40 μm and their resistances should be about 0.1 MΩ when they are filled with Aplysia saline.
2. Pull suction electrodes from polyethylene tubing using a Bunsen burner. Cut a piece of polyethylene tubing about 10 cm long. Hold the tubing on both ends and place it very close to the flame generated by the Bunsen burner while rotating the tube until it becomes soft from the heat. Stretch the tubing carefully along its length while moving it away from the flame. The middle part of the tubing will elongate and narrow as the tubing is pulled.
3. Cut the tubing in half to form two suction electrodes. Suction electrodes are generally applied to the cut ends of nerves or muscles, though they can sometimes be applied en passant.
4. Create hook electrodes for nerve recordings following the protocol described by McManus et al⁸ in sections 3.2-3.13. These electrodes are especially useful when a nerve or muscle is not cut.

3. Hook Electrode Attachment

1. Dissect the animal and remove the buccal mass following the protocol described in McManus et al.⁸ section 4.
2. For recording and stimulation, hook electrodes can be attached to a number of different nerves.
3. To characterize patterns as was done in vivo by Cullins and Chiel⁹, recordings must be obtained from the I2 nerve and muscle that indicates the protraction phase of feeding¹⁶, the radular nerve (RN) that indicates the closure of the food grasper¹⁷, buccal nerve 2 (BN2) and buccal nerve 3 (BN3) that indicate the retraction phase^17,18. Attachment of the hook electrodes follows a procedure similar to that described by McManus et al⁸, section 5.
4. The locations of these nerves refer to the schematic of the Aplysia feeding apparatus shown in Figure 2 of McManus et al⁸. Note that the BN2 trifurcates into branches a, b, and c before going beneath the I1 muscle at the lateral groove. Branch a is the first branch to separate from the main trunk, and is adjacent to the BN3.
5. The nomenclature of branches a, b, and c was used by Warman and Chiel¹⁸. Branches a, b, and c correspond to branches 3, 2, and 1, respectively, in the nomenclature used by Nargeot et al.¹⁹. Furthermore, the RN, the BN1, the BN2, and the BN3 correspond to nerves 1, 6, 5, and 4, respectively, in the nomenclature used by Kandel²⁰ and Scott et al.²¹
6. To study the muscle innervation of the I1/I3 muscle, all the nerves except buccal nerves 2 will be severed from the buccal mass during experiments. Thus, we used a hook electrode to record from the BN2.
7. Since the I2 nerve and the RN will not be attached to the buccal mass, and they are very difficult to access using hook electrodes, it is preferable to apply suction electrodes to record from them instead. We will describe the application of suction electrodes in section 7.
8. Use either a hook electrode or a suction electrode to record from the BN3, because it is easy to access using either kind of electrode. We chose to use a hook electrode for the BN3 recordings to minimize the number of manipulators for holding the suction electrodes, and to save space for other manipulators or equipment.
9. Attach a hook electrode to branch a of BN2 (BN2-a) to initiate rejection-like patterns during experiments. It is useful to attach an additional hook electrode to the BN2-a on the other side, because some neurons respond differently to the ipsilateral vs. contralateral BN2-a stimulation.
10. To help distinguish neurons with unilateral vs. bilateral projections, it is also useful to attach hook electrodes to BN2 and the BN3 on the other side of the buccal ganglia.

4. Ganglia and Muscle Preparation

1. The buccal ganglia, cerebral ganglion and buccal mass will be prepared for the force transducer experiments, in which the cerebral ganglion is attached to the buccal ganglia via the CBCs and the buccal mass is attached to the buccal ganglia via the BN2s only.
2. After attaching the hook electrodes, cut buccal nerve 1 (BN1) and the esophageal nerve (EN) bilaterally, cutting at the attachment point to the buccal mass.
3. Pull the cerebral ganglion forward to move it out of the way of the I2 muscle. Make a cut into the I2 muscle over the radular sac, extend the cut laterally and anteriorly in both directions, and pull the flap of the I2 muscle forward to expose the radular nerve. Cut the two RN branches and make sure that the branches are long enough for suction electrode attachment.
4. Continue the I2 cut in a wide circle around the buccal ganglia, being careful not to cut the BN2s or the BN3s, until the buccal ganglia and the attached part of the I2 muscle can be fully separated from the buccal mass. Cut the bilateral BN3s at the attachment point to the buccal mass, beyond the hook electrode attachment.
5. Apply a thin layer of vacuum grease to the notch in the recording dish described above that connects the back chamber and middle platform, using a pipette tip to pick up a glob of vacuum grease and spread it over the notch.
6. Apply a thin layer of Quick-Gel super glue to the glass bottom of the front chamber where the buccal mass will be placed, just in front of the Sylgard base of the middle platform.
7. Carefully transfer the cerebral ganglion, buccal ganglia and buccal mass to the recording dish (Figure 1) described in section 2, making sure that none of the hook electrodes are pulled tightly, which could damage the nerves.
8. Carefully place the buccal mass on the glue in the front chamber of the recording dish, to ensure that its ventral surface is glued to the bottom of the dish. Be sure to keep the ganglia and electrodes from touching the glue. Add Aplysia saline⁸ (460 mM NaCl, 10 mM KCl, 22 mM MgCl₂, 33 mM MgSO₄, 10 mM CaCl₂, 10 mM glucose, 10 mM MOPS, pH 7.4-7.5) to the dish, which will induce the glue to set.
9. If the dish must be transferred to another microscope for preparing the buccal ganglia for extracellular soma recordings, be very careful with the hook electrodes. Group the electrodes on one side of the buccal mass together, and also group the electrodes on the other side of the buccal mass together. Carefully hold the electrodes by grasping the lab tape that covers the connector pins, again making sure that none of the electrodes are pulled tightly.
10. When the dish is positioned under the microscope, the electrodes should be draped gently over the sides of the dish and rest on the platform next to the dish.
11. During breaks and between stages of the experiment, aerate the saline in the buccal mass chamber using an aquarium airstone.
12. Use forceps to grab the sheath of the cerebral ganglion and pull it into the back chamber, ensuring that the CBCs run through the notch. Pin the cerebral ganglion using nerves other than the CBCs to avoid damage to the intact CBCs.
13. Apply more vacuum grease over the CBCs, and then add more Aplysia saline to both chambers of the dish, so that the ganglia are completely submerged. Ensure that the top of the vacuum grease is slightly higher than the Sylgard wall so that no leakage will occur between the chambers.
14. To stabilize the buccal ganglia, first pin the ends of the BN3s, then the BN1s and the ENs on the Sylgard base of the middle platform (Figure 2). Since the BN3s will be recorded using hook electrodes, the pins should be placed more distally than the attachment points of the hook electrodes.
15. Use two pins, bent 90 degrees, as hooks to stretch and anchor the CBCs, so that the CBCs will not be damaged (Figure 2).
16. Pin down the RN branches between the back chamber and the buccal ganglia. Then the I2 muscle will be on top of the RNs. To expose the I2 nerve, use forceps to grab the I2 muscle and pull it over the buccal ganglia. Pin two corners of the I2 muscle to avoid damage to the I2 nerve.
17. Sever the I2 nerve distal to the point at which its two branches merge into the I2 muscle. Make sure the muscle is still innervated to be comparable to the in vivo recordings. Cut away the rest of the I2 muscle and flip the I2 nerve back and pin it down between the two RN branches (see Figure 2, inset).
18. Adjust the locations of the pins to stretch and add tension if a nerve is too loose, or to release tension if a nerve is too tight. To further stabilize the buccal ganglia, add more pins on the sheath between nerves.
19. Since the buccal ganglia are placed caudal side up, rotate the buccal ganglia if the neurons of interest are on the rostral side. To rotate one of the two buccal ganglia, use fine forceps to grab some excess sheath of the CBC where it is near to the buccal ganglia and pin it down between the BN2 and the BN3. In some ganglia, it may be more convenient to pin it down between CBC and BN3.
20. Add an additional pin on the sheath of the buccal ganglion on the side close to the front chamber to minimize the movement of the buccal ganglion.
21. To trim the sheath covering the buccal ganglia, use fine forceps to grab the sheath on the side close to the back chamber, and then cut away the excess sheath with fine scissors without exposing the cell bodies. In order to minimize damage, only remove the minimum amount of sheath necessary to see the cell bodies.
22. After the sheath of the buccal ganglia is trimmed, pull the I2 nerve and the RNs over the buccal ganglia and pin them down between the buccal ganglia and front chamber to further rotate the buccal ganglia. (See Figure 2).
23. To wash out any remaining magnesium chloride⁸ that was used to anesthetize the animal before dissection, replace the Aplysia saline in the dish with fresh Aplysia saline.

5. Electrically Connecting Hook Electrodes

1. After the ganglia and muscle are prepared, carefully transfer the dish to the vibration isolation table for the experiments.
2. Attach all electrode pins to their sockets on the BNC cables that connect to the amplifiers (A-M Systems model 1700 amplifier). Again, make sure that the electrodes are not pulled tightly while doing this. Make sure that the electrodes are correctly attached to their appropriate cables and that the polarities are correct.

6. Setting up the Extracellular Glass Electrodes for Soma Recordings

1. Fill the electrode with Aplysia saline using a syringe attached to a piece of polyethylene tubing of around 15-20 cm. Attach the free end of the polyethylene tubing to the end of the glass electrode. Pull back on the plunger of the syringe to fill up the electrode with Aplysia saline.
2. Place the filled extracellular glass electrode in the notch of the holder on the manipulator. Use the manipulator to place the electrode tip into the Aplysia saline containing the buccal ganglia.
3. Insert a silver/silver chloride wire soldered to a male gold connector pin into the electrode to serve as the recording wire. Place another silver/silver chloride wire soldered to a male gold connector pin directly into the Aplysia saline within the section of the recording dish containing the buccal ganglia to act as the reference wire. Connect both the recording and reference wires to the BNC cable that connects to the amplifier.
4. If there is enough room for more manipulators, additional extracellular glass electrodes can be added to record multiple neurons simultaneously.

7. Setting up the Suction Electrodes for the Nerve Recordings

1. Trim the narrow end of the suction electrode tip to match the diameter of the nerve. The inner diameter of the electrode tip should be similar to or slightly smaller than the nerve's diameter to ensure tight suction.
2. Since the I2 nerve and the RN are very close to each other, their electrodes can be held by the same manipulator to save space. Place two electrodes in two notches of the same holder. Rotate the two electrodes and ensure that their tips are close to one other. Choose one of them for the I2 nerve recording, the other one for the RN recording.
3. Place the electrode tip in the Aplysia saline within the recording dish containing the buccal ganglia. Attach the free end of the polyethylene tubing on the syringe to the suction electrode. Use the syringe to fill up the electrode with Aplysia saline. Move the electrode tip close to the end of the target nerve, i.e. the I2 nerve, and use the syringe to suck the nerve into the electrode. The length of the nerve within the electrode should be about 0.5-1.0 mm to ensure a tight seal.
4. Repeat the suction for the electrode that will be attached to the RN.
5. Connect the electrodes to the corresponding BNC cables as described in section 6.3.

8. Setting up the Force Transducer to Measure the I1/I3 Muscle Contraction

1. To attach the force transducers to the muscle, use silk sutures. Bend the curved needle of each suture, and tie the suture to the force transducer. Gently grab and lift a small amount of muscle with forceps and, holding the needle in another set of forceps, insert the needle through the muscle up to the bent point in the needle (Figure 1).
2. Transducers can be attached either dorsally or laterally on the I1/I3 muscle. Dorsal attachment allows measurement of contractions evoked by activation of either the left or right side of the muscle. Lateral attachment will show stronger forces for the majority of neurons, but will only allow measurement of contraction on the side to which the transducer is attached.
3. To help identify neurons that may activate the anterior, posterior, or both regions of I1/I3, attach a force transducer to the posterior part of the muscle, just anterior to the pharyngeal tissue, and attach another force transducer to the anterior part of the muscle, at the jaws (Figure 1; note hooks).
4. Lift the force transducers until the sutures are pulled taut, but do not overstretch. To check this, view the measurement from the force transducer when the suture has some slack in it, and then lift the transducer until the measurement is slightly above this baseline level.

9. Identifying Motor Neurons within a Motor Pool

1. This protocol describes a process for extracellularly identifying motor neurons within a motor pool. We used AxoGraph software to monitor the activity of individual neurons, multiple nerves, and the muscle (EMG signal or contraction forces). In this protocol, we used the contraction forces of the muscle as an illustration for the process of identifying motor neurons; in other experiments, we used EMG as well, and the setup for such experiments is very similar (see Discussion).
2. To locate a candidate neuron, use the manipulator to gently press the tip of the extracellular glass electrode down onto the sheath over the center of the neuron soma⁵ (Figure 3), which is the best location for stimulation and recording selectivity⁵. Since the threshold current for activating a neuron increases linearly with electrode-to-soma distance⁵, the stimulation selectivity will become worse when the electrode is moved away from the center of the target neuron toward a neighboring neuron.
3. To identify a motor neuron, first directly stimulate the neuron using the extracellular glass electrode to ensure that only this neuron is firing, and examine whether it innervates the muscle. Then, extracellularly record from this neuron to establish a one-for-one relationship between the extracellular soma recording and the nerve recordings, which is also crucial for neuron identification.
4. Since most extracellular amplifiers do not allow simultaneous stimulation and recording in a channel, set the channel used to stimulate and record the soma (the soma channel) to stimulation mode and apply a brief anodic current (e.g. 6 msec for Aplysia neurons⁵) to the soma (Figures 4A, 5A; note arrows 1 in both figures), starting from a low current (e.g. 200 μA), and gradually increasing the current until the neuron bursts.
5. Once the neuron is activated to burst, one should immediately switch the soma channel from the stimulation mode to the recording mode (Figures 4A, 5A; note arrows 2 in both figures). However, there will still be delays between the soma stimulation and recording because of human response delays.
6. If the neuron fires for a reasonable amount of time, it should be possible to observe one-for-one corresponding action potentials from the soma recording and on the nerve(s) through which the neuron projects (Figures 4B, 5B; note dashed lines), as well as the forces generated by the neuron (Figures 4A, 5A; note blue boxes). If the neuron stops firing before the soma recording begins, increase the current to activate it for a longer time.
7. The signal-to-noise ratio of the extracellular recordings depends on the electrode location and soma size. The extracellular recording will become bigger as the soma size increase and electrode-to-soma distance decreases. Since the noise only varies in a narrow range, the signal-to-noise ratio will also increase as the soma size increases and the electrode-to-soma distance decreases. The most common range of the signal-to-noise ratio is from 4:1 to 8:1.
8. The motor neurons can be identified based on their characteristics, such as soma location, nerve projection and muscle innervation^4,6,7. Since only the two BN2s are attached to the buccal mass, by monitoring the activity on the BN2s, one can ensure that the muscle contraction force is only caused by the neuron activated by the extracellular stimulation.
9. For example, B3 is a large motor neuron for the I1/I3 muscle (300-400 μm in soma diameter in animals weighing 200 to 350 grams), located on the rostral side of the buccal ganglion (Figure 3). It only projects through the ipsilateral BN2, and innervates both the anterior and posterior parts of the I1/I3 muscle. Most of the time, activating it generates a larger anterior than posterior force (8 out of 9 experiments).
10. After a neuron is identified, its activity can be recorded in different feeding-like behaviors via the extracellular glass electrode (Figures 4 and 5), which can be elicited as described below. The extracellular recording on the soma will be much more specific than the nerve recordings, which include the activity of many different neurons.
11. To induce egestive-like motor programs, stimulate BN2-a with 1-2 min of pulses¹⁹ (2Hz, each pulse is 1 msec). This stimulation reliably generates egestive patterns in this setting. With sufficient current (e.g. 300 μA), patterns may persist for the duration of the stimulation. Sometimes there will be one more pattern that occurs shortly after the stimulation ends.
12. To induce ingestive-like motor programs, place a few crystals of solid carbachol directly onto the sheath of the cerebral ganglion¹⁵. If one wants to control the level of carbachol exposure, use a solution of 1 to 10 mM carbachol in Aplysia saline. Higher concentrations are more likely to induce responses. Repetitive patterns generally begin within five minutes, and may last for approximately ten to fifteen minutes before beginning to run down.
13. After washing out carbachol several times and waiting for at least 30 min, a subsequent application of carbachol can be added to the cerebral ganglion to induce more ingestive-like motor patterns.
14. After multiple motor neurons for the particular motor pool have been identified and characterized during motor programs, one may develop a greatly simplified diagnostic method that requires minimal information for quickly identifying these neurons in future work (Figure 6), e.g. in the suspended buccal mass preparations or in vivo. The criteria may include soma size and location, nerve projection, unit size on the nerves, and timing of activity during motor patterns.

Results

Figures 4 and 5 show typical results used to identify two I1/I3 motor neurons. Figure 4 shows the soma recordings of a large motor neuron, B3, during egestive-like and ingestive-like patterns (Figures 4C, 4D). The one-for-one corresponding spikes on the soma channel and the ipsilateral BN2 channel (Figure 4E) show that the specificity of B3 soma recording was maintained during patterns. B3 fires during the middle-to-late retraction phase...

Discussion

In animals with large identified neurons, such as mollusks (for example, Lymnaea, Helix, and Aplysia), analysis of motor pools is typically done using intracellular recording^1,2,3,4. In this protocol, we describe a process for uniquely identifying the motor neurons for a motor pool using an extracellular technique. We used the force measurements as an illustration of this process. One could also use EMG to measure muscle innervations. Briefly, to do so, the protocol needs to be ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Sodium chloride	Fisher Scientific	S671	Biological, Certified
Potassium chloride	Fisher Scientific	P217	Certified ACS
Magnesium chloride hexahydrate	Acros Organics	19753	99%
Magnesium sulfate heptahydrate	Fisher Scientific	M63	Certified ACS
Calcium chloride dihydrate	Fisher Scientifc	C79	Certified ACS
Glucose (dextrose)	Sigma-Aldrich	G7528	BioXtra
MOPS buffer	Acros Organics	17263	99%
Carbachol	Acros Organics	10824	99%
Sodium hydroxide	Fisher Scientific	SS255	Certified
Hydrochloric acid	Fisher Scientific	SA49	Certified
Single-barreled capillary glass	A-M Systems	6150
Flaming-Brown micropipette puller model P-80/PC	Sutter Instruments		Filament used: FT345B
Enamel coated stainless steel wire	California Fine Wire		0.001D, coating h
Household Silicone II Glue	GE
Duro Quick-Gel superglue	Henkel corp.
A-M Systems model 1700 amplifier	A-M Systems		Filter settings: 10-500 Hz for the I2 nerve/muscle; 300-500 Hz for all the other nerves
Pulsemaster Multi-Channel Stimulator	World Precision Instruments	A300
Stimulus Isolator	World Precision Instruments	A360
AxoGraph X	AxoGraph Scientific		Software for recordings
Gold Connector Pins	Bulgin	SA3148/1
Gold Connector Sockets	Bulgin	SA3149/1
Sylgard 184 Silicone Elastomer	Dow Corning
100 x 15 mm Crystalizing Dish	Pyrex
High Vacuum Grease	Dow Corning
Pipet Tips	Fisher Scientific	21-375D
Minutien Pins	Fine Science Tools	26002-10
Modeling Clay	Sargent Art	22-4400
Whisper Air Pump	Tetra	77849
Aquarium Tubing	Eheim	7783	12/16 mm
Elite Airstone	Hagen	A962
Vannas Spring Scissors	Fine Science Tools	15000-08
Dumont #5 Fine Forceps	Fine Science Tools	11254-20
Kimwipes	Kimberly-Clark	34155