Method for the Assessment of Neuromuscular Integrity And

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Journal of Neuroscience Methods 264 (2016) 40–46

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Journal of Neuroscience Methods

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Short communication

Method for the assessment of neuromuscular integrity

and burrowing choice in vermiform animals

∗

C. Bainbridge, A. Schuler, A.G. Vidal-Gadea

School of Biological Sciences, Illinois State University, 339 Science Laboratory Building, Normal, IL 61790-4120, USA

h i g h l i g h t s

•

Detailed instructions for the construction of burrowing assays are provided.

•

Assays permit the study of the effect of muscular exertion on locomotion.

•

Assays are useful to study burrowing preference including magnetic orientation.

a r t i c l e i n f o a b s t r a c t

Article history: Background: The study of locomotion in vermiform animals has largely been restricted to animals crawling

Received 31 December 2015

on agar surfaces. While this has been fruitful in the study of neuronal basis of disease and behavior, the

Received in revised form 26 February 2016

reduced physical challenge posed by these environments has prevented these organisms from being

Accepted 29 February 2016

equally successful in the study of neuromuscular diseases. Our burrowing assay allowed us to study the

Available online 3 March 2016

effects of muscular exertion on locomotion and muscle degeneration during disease (Beron et al., 2015),

as well as the natural burrowing preference of diverse Caenorhabditis elegans strains (Vidal-Gadea et al., Keywords: 2015).

Burrowing

New method: We describe a simple, rapid, and affordable set of assays to study the burrowing behavior

C. elegans

Locomotion of nematodes and other vermiform organisms which permits the titration of muscular exertion in test

animals.

Results: We show that our burrowing assay design is versatile and can be adapted for use in widely

different experimental paradigms.

Comparison with existing method(s): Previous assays for the study of neuromuscular integrity in nematodes

relied on movement through facile and homogeneous environments. The ability of modulating substrate

density allows our burrowing assay to be used to separate animal populations where muscular ﬁtness or

health are not visible differentiable by standard techniques.

Conclusion: The simplicity, versatility, and potential for greatly facilitating the study of previously chal-

lenging neuromuscular disorders makes this assay a valuable addition that overcomes many of the

limitations inherent to traditional behavioral tests of vermiform locomotion.

1. Introduction 2012). However, much progress remains to be made in order to

understand this behavior. Most work to date focused on poly-

Burrowing is one of the most widespread forms of animal loco- chaetes and annelids which are behaviorally accessible but not

motion. Many organisms use burrowing as their primary form of genetically tractable (Topoliantz and Ponge, 2003; Dorgan, 2015).

locomotion through a wide range of substrates. For example, ter- Unlike other forms of locomotion (e.g. swimming, crawling; Vidal-

restrial vertebrates like the sandﬁsh lizard (Maladen et al., 2009) Gadea et al., 2011) burrowing takes place through environments

and various species of burrowing snakes (Marin et al., 2013) use that are often heterogeneous and therefore pose varying demands

burrowing through a wide range of substrates. Burrowing is also on the musculature of burrowing organisms. For example, work

frequently seen in subsea terrains (e.g. razor clams Winters et al., on burrowing on clams characterized the immense muscular effort

required to burrow through heterogenous environments (Winters

et al., 2012). The variable demands posed on the organism makes

∗ burrowing especially well-suited for the study of neuromuscular

Corresponding author. Tel.: +1 309 438 5220; fax: +1 309 438 3722.

E-mail address: [email protected] (A.G. Vidal-Gadea). disorders where it can be crucial to discern the functional limits

http://dx.doi.org/10.1016/j.jneumeth.2016.02.023

C. Bainbridge et al. / Journal of Neuroscience Methods 264 (2016) 40–46 41

of healthy and diseased musculature. The use of burrowing assays effects of varying levels of muscular exertion on muscle health and

allows experimenters to ﬁnely titrate substrate density, thus con- locomotion. This is accomplished by constructing pipettes ﬁlled

trolling the amount of muscular exertion required of test animals. with regular NGM (rather than chemotaxis) agar, and that have

Recently, we developed a burrowing assay which permitted us OP50 bacterial food at regular intervals to prevent starvation. (3)

for the ﬁrst time to conduct genetic screens that separate healthy Assessing burrowing choice involves injecting animals into the cen-

animals from those with dystrophic muscles (Beron et al., 2015). ter of pipettes ﬁlled with chemotaxis agar and equidistant from test

This work restricted itself to illustrating the usefulness of this and control stimuli at opposite ends of the pipette. (4) To ﬁlm and

new assay without providing essential directions for its success- quantify the kinematics of burrowing behavior we replace 5 mL

ful performance, or adaptability to additional approaches. Here we plastic serological pipettes with 2 mL borosilicate glass Pasteur

present detailed step by step directions for the successful perfor- transfer pipettes which are smaller in diameter and more transpar-

mance of this assay to challenge and assess muscular integrity. ent than their plastic counterparts. This permits the clear ﬁlming

These burrowing assays offer a reliable way to assess neuromuscu- of burrowing animals under different experimental conditions.

lar degeneration by using cost effective and easy to use materials

that are equally amenable to the performance of genetic screens, as

3. Results and discussion

well as teaching activities. We discuss the potential for combining

this approach with current force measurement approaches.

3.1. Agar preparation for burrowing assay

Recently, we discovered that nematodes such as Caenorhabdi-

tis elegans detect and orient to earth-strength magnetic ﬁelds. We

The preparation of agar for burrowing studies requires some

adapted a T-maze assay in order to present burrowing worms with

considerations unique to these studies and differing depending on

two choices and went on to show that worms use the Earth’s mag-

the type of assay to be conducted. For short term choice assays

netic ﬁeld to guide vertical migrations in a satiation dependent

where animals will not be tested over long periods of time (under

manner (Vidal-Gadea et al., 2015). We describe a simple and novel

12 h) a standard chemotaxis agar recipe can be used (Hart, 2006)

set of assays designed to facilitate the study burrowing in C. elegans

which avoids the use of salts which might otherwise interfere with

and other vermiform animals. We provide detailed instructions

the tested orientation behavior of worms. Brieﬂy, this consists of 3%

in the setup and execution of burrowing assays designed to test

agar, 5 mM KPO [pH6], 1 mM CaCl , and 1 mM MgSO which are

muscular integrity, study burrowing kinematics, and determine 4 2 4

autoclaved for 20 min, poured into 10 cm Petri plates, and allowed

burrowing preference under differential burrowing conditions.

to dry overnight. The amount of agar used can be modiﬁed to obtain

Harnessing burrowing behavior of nematodes and other vermi-

agar of different densities. For long term assays, where worms need

form animals will provide a wealth of new insights into the study

to survive in plates for a day or longer, we used standard nema-

of many neuromuscular diseases as well as the understanding of

tode growth media agar (Brenner, 1974), as this agar contains all

animal behavior.

the salts and nutrients required for worms to grow and thrive.

Brieﬂy, 1 L of NGM agar is made by autoclaving of 3 g NaCl, 17 g

2. Materials and methods

agar, and 2.5 g peptone in 975 mL of dH2O. After the solution cools

◦

to 55 C 25 mL of KPO4, 1 mL 1 M MgSO4, 1 mL 1 M CaCl2, and 1 mL

2.1. Culturing and preparation of C. elegans for burrowing assays

of 5 mg/mL cholesterol (in ethanol) are added, in that order, to pre-

vent salt precipitation at intermediate pHs. The use of NGM agar

Wild-type C. elegans were raised on nematode growth media

during long term assays is necessary as worms need cholesterol

(NGM) agar plates seeded with OP50 strain bacterial lawns for food

and salts for prolonged survival and would not develop normally

as previously described (Brenner, 1974). Animals were grown at

◦ in chemotaxis agar as it lacks salts and cholesterol. Because worms

20 C, in a 37% humidity room, in infection-free plates and never

will be burrowing through the agar, it is imperative that it be free

allowed to starve or to overpopulate their plates before testing.

of salt crystals as these have the potential to harm worms as they

Burrowing assays are carried out on day 1 adult animals. Animal

move between them. Therefore the pH of each solution used in

populations were synchronized by bleaching adult hermaphrodites

the agar preparation must be carefully monitored and each pipette

(Porta-de-la-Riva et al., 2012) and placing isolated eggs into agar

must be inspected before and during use. It is particularly impor-

plates. Worms have been described to rapidly accumulate random

tant to add the CaCl2 last as calcium may fall out of solution if

mutations (Denver et al., 2004). Behavioral parameters such as

added prematurely. Air bubbles within the pipette must be absent

locomotion are susceptible to mutations in many genes and will

as these allow worms to exit the agar and alternate between bur-

show signiﬁcant changes over many generations (personal obser-

rowing and crawling. Air bubbles can have different causes and

vation). Therefore to prevent unwanted mutations from affecting

may be introduced during the preparation of the agar. To prevent

our results we instituted a lab practice whereby every strain

these type of air bubbles it is important to cover the glass con-

being worked with is replaced (from frozen stocks) every three

tainer with the cooling agar with plastic wrap (Fig. 1A). As the agar

months.

cools in the container, the hot air within the sealed container also

cools decreasing in volume following the ideal gas law (PV = nRT).

This results in the creation of a vacuum that pulls air out of solu-

2.2. Burrowing pipette preparation

tion resulting in homogenous (bubble-free) agar. A second source

of air inside agar pipettes is time. Agar pipettes may be stored

Burrowing assays are conducted in 5 mL serological or 2 mL

◦

in a 4 C refrigerator for short periods of time but must be used

borosilicate glass pipettes (ExactaCruz, CA) ﬁlled with chemotaxis

within one or two weeks from their construction (ideally no later

agar (Hart, 2006). The construction of the pipettes differs depending

than two or three days). Eventually, agar within the pipettes will

on the intended parameter being evaluated. We have devised four

dry and shrink, creating air pockets over which worms are able to

different conﬁgurations which we use to study diverse phenomena:

crawl freely. Similarly, an additional source of air pockets are the

(1) Different density agars can be used to present animals with

holes bored into the pipettes to accept chemicals or animals. These

varying muscular challenges. Animals are then raced while they

holes need to be made as close to the start of the assay as possi-

burrow toward an attractant in pipettes containing either different

ble to prevent agar from drying and introducing air pockets in the

environmental conditions, or different worm strains (Fig. 1). (2) A

pipettes.

second pipette conﬁguration permits the study of the long term

42 C. Bainbridge et al. / Journal of Neuroscience Methods 264 (2016) 40–46

◦

Fig. 1. Preparation of burrowing assay. (A) Use a microwave oven to boil Chemotaxis agar. (B) When the agar cools to ∼50 C, use a pipette controller to draw 5 mL into

a plastic serological pipette. (C) Temporarily plug the tapered end of the pipette with paraﬁlm prior to removing the pipette controller. Seal both ends of the pipette with

paraﬁlm. (D) After the agar solidiﬁes, use a heated blade to score and snap both ends of the pipette. This will ensure consistent pipette length, diameter, and the remove air

pockets that would otherwise be left at the top of the pipette during ﬁlling. Seal new ends with paraﬁlm to prevent formation of humidity gradients or air bubbles. (E) Heat

a 5 mm metal rod and gently melt a hole in the pipette 1 cm in from one of its ends to create a well for an attractant or bacteria food. (F) Concentrated 1.5 mL of OP50 E. coli

culture by centrifugation at 7000 × G for about two minutes. (G) After removing all but 50 ␮L of supernatant, resuspend bacteria and inject 25 ␮L into the well. Seal the cavity

using Paraﬁlm. (H) Heat a 2 mm metal rod and gently create a hole in the pipette 1 cm in from the opposite end of the pipette to make the animal injection site. Make sure

that the hole only reaches the center of the pipette and that no plastic debris is left after the creation of either hole (I) Place a 3 ␮L drop of liquid nematode growth media

buffer (NGM) onto a paraﬁlm strip. (J) Use a platinum wire to transfer between 20 and 50 worms from a culture plate into the NGM droplet. (K) Worms should be readily

seen swimming in the water drop. Inset: Remove small amount (∼2 ␮L) of NGM to concentrate worms prior to transferring them into the injection site. (L) Draw the worms

into a glass capillary tube using a Drummond micropipette. (M) Gently stab injection site with the capillary tip while expelling the worms directly into the agar. (N) Use a

stereo microscope to conﬁrm that worms are burrowing in the agar and not still swimming in the liquid NGM. (O) Representative scoring of burrowing pipette assays. White

lines mark the 1 cm and 4 cm cut-offs for assay scoring. Red dots indicate animal position. (P) Graph indicating the fraction of animals that migrated more than 4 cm from

the animal injection site. (For interpretation of the references to color in this ﬁgure legend, the reader is referred to the web version of this article.)

3.2. Filling of pipettes reheated in a microwave and Saran Wrap was used to remove air

bubbles accidentally introduced. With the pipette controller still

Polystyrene serological pipettes (5 mL) were ﬁlled with chemo- attached, the tapered end of the pipette was temporarily sealed

taxis agar by drawing agar from the bottom of a 1 L ﬂeaker using an using a piece of paraﬁlm folded several times to clog the tip of the

Accu-jet pro pipette controller (Brandtech, CT). NGM agar can be pipette. Applying pressure to the tapered end (to prevent leaking),

substituted for long term burrowing where the worms will develop the pipette controller was removed and a piece of paraﬁlm strip was

and will require nutritional supplements from the agar. This is used to seal this end of the pipette (Fig. 1C). The pipette was then

done to control for muscle deterioration caused by lack of nutri- placed vertically in a container to cool at room temperature. After

tion rather than exertion. Prior to ﬁlling the agar was allowed to about ten minutes the agar in the pipettes reached room temper-

◦

cool to roughly 50 C (or until cool enough to permit your palm to ature and became solidiﬁed. This protocol allowed us to construct

contact the container for ﬁve seconds with no pain). Pipettes were pipettes ﬁlled with agar ranging between 0.5% and 9%. In our experi-

slowly ﬁlled just below the safety cotton plug taking care that no ence it was challenging to obtain bubble-free agar of homogeneous

air bubbles were introduced (Fig. 1B). If necessary, the agar was consistency outside of this range.

C. Bainbridge et al. / Journal of Neuroscience Methods 264 (2016) 40–46 43

3.3. Resizing of pipettes 3.4.3. Placement of anesthetic

During our choice assays worms were injected in the middle

Filling the pipettes with agar leaves an air pocket above the cot- of the pipettes and test or control solutions were introduced in

ton guard that if left unattended will result in the top end of the equidistant holes at each end of the pipettes. To be able to tally

pipette drying and creating unintended humidity and density gra- worms once they made a choice, we introduced 2 ␮L of 1 M sodium

dients. In order to prevent this the ends of the pipettes must be azide (NaN3) into these holes just prior to the beginning of the assay.

removed as soon as the agar has ﬁnished solidifying. To do this we This timing was important as it prevented the anesthetic from dif-

used vise-grip pliers to hold a utility knife blade (Alvin, CT) over fusing too far and effectively preventing the worms from making

a bunsen burner until red hot. Pipettes were aligned and the hot their behavioral choice. We used the geomagnetotactic burrowing

utility blade was used to score the pipettes taking care that the cut behavior of C. elegans to exemplify this type of assay. This is based

was over agar ﬁlled sections of the pipettes (to remove air pockets), on our previous ﬁnding that well fed and starved worms burrow

and that all resulting pipettes were of the same size (Fig. 1D). After upwards or downwards (respectively) when injected into vertically

scoring the pipettes could be easily snapped and we once again aligned pipettes (Vidal-Gadea et al., 2015). However, the behavioral

used paraﬁlm to thoroughly seal the new (exposed) ends. Depend- choice assay can be used to assess preference of test or control sub-

ing of the intended use pipettes can be used as is, which roughly stances as well. This is done by placing these solutions at the ends

yields 24 cm of burrowing pipette. Alternatively, for use in quick of the pipettes two days before testing so that a chemical gradient

races pipettes can be halved resulting in two 12 cm assay tubes for had a chance to develop across the pipette.

each pipette prepared. We primarily used whole pipettes when per-

∼

forming multi-day assays and half pipette tubes for shorter ( 2 h) 3.5. Collection of worms for assays

burrowing races.

3.5.1. Collection of worms for assays by picking

Worms were obtained from well-fed plates by individually

3.4. Insertion and stimuli site preparation transferring animals with the help of a sterilized platinum wire.

Whenever possible animals that were crawling outside the E. coli

3.4.1. Preparation of perforations lawn were chosen as this minimized the amount of bacteria inad-

Injection sites were made using the back of a heated 2 mm drill vertently transferred along with them. Worms were then released

bit taking care that the hole was perpendicular to the surface of the into a 3 ␮L droplet of liquid NGM buffer atop a strip of paraﬁlm

pipette, and that it stopped short of the center of the tubes. Care was (Fig. 1J). About 50 worms were thus transferred and allowed to

taken to ensure that there was no melted plastic was left coating the swim in the liquid. This also removed bacteria potentially coat-

interior of the hole as this would prevent the injection of animals ing them. We used a glass capillary mounted on a Drummond

into the agar. Each hole was therefore inspected under a micro- micropipette to concentrate the worms in the droplet by remov-

scope and any plastic found was removed with forceps. Holes for ing 2 ␮L of the 3 ␮L of liquid NGM (Fig. 1K). This minimized the

choice assays were placed at the center of the pipettes while holes amount of liquid injected along with the worms and it is necessary

for races or long term incubation were placed 1 cm away from one because the animals are unable to overcome the surface tension of

end of the pipettes (Fig. 1H). In addition to making holes to insert water and would remain permanently suspended in solution if the

worms we also made holes to place attractants, anesthetics, or OP50 droplet is too large and is not readily absorbed by the surrounding

bacteria food depending on the assay. We followed a similar proto- agar. By using a Captrol III micropipette (Drummond, AL) a glass

col described above but used a 5 mm drill bit to make a larger hole capillary can easily be used to draw most of the liquid NGM in the

␮

in order to accommodate a larger volume (up to the 25 L of OP50 droplet. We then used the micropipette to transfer 1 ␮L of worms

we inject for races and long term assays). The volume of these hole in NMG into the assay pipettes (Fig. 1L). An alternative to remov-

could be doubled if necessary by increasing the depth of diameter ing small volumes of NGM prior to injection would be to inject the

of the holes. We made holes (two for the choice assay, one for the whole volume and remove excess liquid NGM from the assay under

race assays) 1 cm away from the end of the pipettes (Fig. 1E). As a dissecting scope with a Kimwipe. The authors do not recommend

before, care was taken that no plastic remained coating the inner relying on this method since it often results in a large loss of worms.

surface of the perforation. Paraﬁlm was used to temporarily seal

the holes which protected them from desiccation up to overnight. 3.5.2. Collection of worms for assays by centrifuge

For some assays satiation is an important parameter. In our mag-

netic assays the time between worm removal (from its home plate)

3.4.2. Preparation and placement of OP50 food

and the beginning of the assay must be kept to within 5–10 min.

We used OP50 bacteria both as a powerful attractant during our

When time is of the essence it is possible to use 1.5 mL of liquid

burrowing races, and as a food source during our long term burrow-

NGM to quickly remove a large number of worms from their home

ing assays. OP50 Escherichia coli was cultured in LB broth overnight

plate. When doing this the liquid NGM must be gently deposited in

◦

in a 37 C shaker. The bacteria were concentrated by placing 1.5 mL

the culture plate to avoid bringing OP50 bacteria into solution. The

of OP50 in eppendorf tubes and centrifuging them at 7000 × G for

plates are gently tapped to allow the worms (but not the bacteria)

2 min. All but 50 ␮L of the supernatant were discarded and the bac-

to enter solution. The plates are then tilted and the liquid with the

terial pellet resuspended in this volume by serial pipetting (Fig. 1F).

worms is pipetted into a 1.5 mL eppendorf tube where worms can

For our burrowing races we placed 25 ␮L of this concentrate in the

be partially pelleted by centrifugation at 7000 × G. Worms are not

pipettes, sealed the hole with paraﬁlm and incubated the pipettes

much denser than water so they never pellet very well, however

◦

in the 4 C refrigerator for two days to allow the formation of a

this method will allow to concentrate many worms in the bottom of

chemical gradient across the length of the pipettes (Fig. 1G). For

a tube where they can be pipetted out into an agar pipette relatively

our long term burrowing assays we placed holes 5 cm apart in quickly.

the pipettes and used 25 ␮L of concentrated OP50 to ensure that

worms had enough food to thrive, but also that they had to burrow

3.6. Injection of worms into pipettes

across the pipette in order to keep ﬁnding food. We incubated the

◦

pipettes in the 4 C refrigerator for two days following the addition

Paramount for the success of burrowing assays is that worms

of concentrated OP50 to establish chemical gradients in the agar.

be deposited directly into the agar rather than remain trapped in a

44 C. Bainbridge et al. / Journal of Neuroscience Methods 264 (2016) 40–46

drop of liquid NGM. This is the case because worms are normally between 1.14 and 0.68 cm/min, however, over 22 strains across

unable to break the surface tension of a drop of water and therefore four physiological conditions (>1500 animals) show a variability

remain trapped within it as long as the drop persists. This has the of 0.21 ± 0.04 cm/min (mean/stdev; Vidal-Gadea et al., 2011). Sim-

unintended consequence of forcing worms to swim for prolonged ilarly, while the burrowing velocity of animals with diverse genetic

periods of time, a behavior which unlike burrowing, is character- backgrounds ranges between 0.05 and 0.46 cm/h the variability for

ized by fast frequency and low amplitude. Because the agar in the each strain performing this behavior was 0.05 ± 0.03 cm/h (Beron

pipettes tends to be fresh, it is considerably humid, thus prolonging et al., 2015).

the time it takes for it to absorb the NGM droplet and release the

worms. We have observed worms still swimming at their injection

3.7.2. Data collection for long term burrowing assays

site several days after being injected. Finally, because during races

After worms have been allowed to burrow in agar pipettes for

and during choice assays worms are performing a time-sensitive

several days they must be retrieved in order to assess their motor

task, it is crucial that all animals enter the test at the same time. For

skills by ﬁlming and tracking populations of crawling animals.

these reasons it is important to inject worms into the agar under a

Alternatively their musculature can be evaluated by performing

dissecting microscope to ensure that they are successfully released

confocal microscopy of genetically-tagged or immunohistochem-

inside the agar (Fig. 1N). One way to make sure that worms are

ically treated animals. These tests require that animals be safely

released into the agar is to make sure that there is a small air bub-

removed from the agar pipettes. We removed animals from their

ble in front of the NGM droplet with the worms inside the capillary.

pipettes by using a hot utility blade to score the surface of the

The capillary is then used to stab the agar while pushing the air into

pipettes at regular 3 cm intervals. carefully snapping the pipette

the agar. The tip of the capillary must be well within the agar and

sections allowed the retrieval of sections of agar that were then

away from the walls of the pipette. Once the air has been forced into

placed in a 10 cm petri dish containing a thin layer (5 mL) of chemo-

the agar, the capillary is gently and slowly removed while pushing

taxis agar and liquid NGM (10 mL). The agar sections were further

the worms into the agar. A vacuum forms in the agar which helps

broken into 1 cm slices and worms were retrieved from the liq-

draw the worms into it and places them inside the agar and away

uid NGM. Once a burrowing worm enters the NGM liquid it can

from the NGM solution. This method takes a little practice but it

no longer exit it. This permits its safe retrieval. Because our assay

quickly becomes a sure way to transfer worms from liquid into

seeks to quantify muscular and motor degeneration it is important

solid agar. If worms are not injected into the agar but rather sit in

to remove all animals observed regardless of their apparent health.

a droplet inside the hole it is possible to use a piece of Kimwipe

Animals are then transferred into a recovery agar plate with OP50

to draw excess liquid. However this method has some drawbacks

food awaiting their behavioral or anatomical assessment.

as the worms are only released on the surface of the agar (rather

than within it) severely affecting the timing of the assay. Second,

3.7.3. Data collection from choice assays

most of the worms will be also drawn up with the Kimwipe, reduc-

Following a choice assay worms were paralyzed by the anes-

ing dramatically the number of animals that enter the assay. For

thetic (NaN3) at one of two possible exit points. Worms were

these reasons placing the worms into the agar directly is by far the

then tallied as described above. We then calculate their index of

preferred method.

preference using the formula: MI = (T − C)/(T + C) where MI is the

migration index (e.g. chemotaxis index, magnetotaxis index, etc.),

3.7. Data collection

T is the number of worms that migrated toward the test stimu-

lus, and C the number of worms that migrated toward the control

3.7.1. Data collection for burrowing races

stimulus. Here we exempliﬁed a burrowing choice assay using the

After worms were allowed to burrow toward an attractant for

geomagnetotactic preference of worms (Fig. 2). Because magneto-

2 or 3 h (depending on the assay parameters), the pipettes were

tactic behavior in worms is relatively fragile behavior, the authors

placed under a dissecting microscope and worms were observed

recommend a sample size of >20 assays each with 20–30 animals.

and tallied under dark ﬁeld illumination. A permanent marker was

used to place a dot on the surface of the pipette above each observed

3.8. Use of borosilicate transfer pipettes for ﬁlming of burrowing

worm. This permitted recording the positions of many animals

animals

in several pipettes pending future tallying (Fig. 1O). Once all the

worms in all the pipettes were so marked, two lines were drawn

Some experiments require the ﬁlming of burrowing animals for

1 cm and 4 cm away from the injection sites. This allowed us to

later kinematic analysis. In this type of assays plastic serological

count the number of worms that enter the assay (by virtue of bur-

pipettes are ill suited as their diameter and opacity prevent reliable

rowing) but failed to make progress (those between the 1 cm and

imaging of behaving animals. In these circumstances we replace

the 4 cm marks, and those that successfully burrowed past the 4 cm

plastic serological pipettes with borosilicate glass Pasteur pipettes

mark). This threshold was useful because it removed worms from

instead. Glass pipettes hold 2 mL of agar. Their reduced diameter

the tally that remained at the injection site perhaps dead, or trapped

and increased transparency make them ideal for ﬁlming burrowing

in liquid, thus reducing variability. We therefore calculated the frac-

worms (Supplemental video 1). These pipettes cannot be easily cut

tion of worms that burrowed >4 cm by dividing the number of these

so attractants must be injected into their tapered end, while worms

animals by the number of worms that burrowed >1 cm (Fig. 1P).

are injected into the opposite end of the pipette.

We ﬁnd that in our hands assays consisting of about 50

animals yield best results. This insures that a large number of

animals participate in the assays while maintaining the density 3.9. Compatibility with force measuring techniques

of worms in the pipette low enough to where most worms are

in seldom contact with each other. We typically run at least One of the main usefulness of the burrowing assay presented

twenty assays for each experimental condition but ﬁnd that assay here is its ability to force animals to generate different levels of

variability remains constant after about ten replicates. Result vari- muscle exertion during locomotion. It follows that it may be desir-

ability depends on the strain and experimental condition but is able to combine these assays with force measuring techniques to

largely at par with the variability observed for other described quantify the muscular output of animals. Several techniques are

behaviors. Crawling worms carrying diverse mutations and under presently used to infer force exerted by the muscles of burrow-

different physiological conditions move at a velocity ranging ing animals such as photoelastic measurements (Dorgan et al.,

C. Bainbridge et al. / Journal of Neuroscience Methods 264 (2016) 40–46 45

Fig. 2. Example of burrowing choice assay: Magnetic orientation. (A) Prepare the burrowing choice pipettes by injecting anesthetic into holes at the ends of the pipette and

the worms in the injection site at the center of the pipette. It is important to make sure that worms are injected straight into agar to not bias their burrowing preference.

(B) Once the pipette injections are complete, they can be bundled together and placed into a grounded copper mesh ﬁber cloth sleeve which will act as a faraday cage for

our magnetic burrowing choice assay. The grounded copper cloth should be completely wrapped around the pipettes in order to adequately shield the assays from electric

fields. (C) Orient the shielded pipettes in the direction of the magnetic field. In the case of Earth’s magnetic field we line the pipettes with the field using a compass, and at

◦ ◦

adjust the angle of the pipette to the inclination of the field (approximately 68 along the direction of the field or 112 against the direction of the field) using a clamp. Allow

the assay to run until the animals are paralyzed (approximately 1 h). Once the assay is ﬁnished the copper cloth faraday cage can be unwrapped and the pipettes can be

individually scored (D) Results from the burrowing choice assay are shown. Graph indicates burrowing preference in the presence of a magnetic ﬁeld is satiation dependent.

The magnetic index is calculated as MI = (T − C)/(T + C) where MI is the magnetic index, T and C are the number of animals that migrated to the test stimulus and control

stimulus, respectively. For this conﬁguration of burrowing choice assay a positive MI means animal migration up the pipette, and a negative MI means movement down the

pipette. (E) Representative scoring of burrowing choice assay. In this case the end of the pipette that was oriented upward is indicated with a black arrow (arrow was drawn

on the pipette before running the assay). Animal positions were scored in red marker. (For interpretation of the references to color in this ﬁgure legend, the reader is referred

to the web version of this article.)

2007; Mirbagheri et al., 2015), or calcium activity reporters (Pierce- Appendix A. Supplementary data

Shimomura et al., 2008). While these techniques should be both

applicable to our assay, we feel that microﬂuidic chips probably Supplementary data associated with this article can be found, in

deliver a more controlled environment for these kind of measure- the online version, at http://dx.doi.org/10.1016/j.jneumeth.2016.

ments. We ﬁnd that our assay’s strength lies in its high throughput, 02.023.

its affordability, and its ease which permits its use in a classroom setting.

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