Optomotor Drum

Background

The optomotor reflex is an animal’s tendency to stabilise themselves relative to visual information. We can use this to our advantage in simple behavioural experiments to work out the limits of an animal’s visual capabilities, such as its acuity or ability to see colour contrasts.

We have developed a customisable low-cost optomotor apparatus (AKA “optomotor drum”) , which can be built from readily accessible materials and 3D printed parts. For further details of the background and scope of this resource see our publication: Caves, E.M., Troscianko, J. and Kelley, L.A. (in press) A customizable, low-cost optomotor apparatus: a powerful tool for behaviourally measuring visual capability, Methods in Ecology and Evolution. DOI: 10.1111/2041-210X.13449

This video guide takes you through the entire assembly process:

Worked Example

Here we illustrate the process in detail of constructing and programming the optomotor apparatus, creating stimuli, choosing parameters for the experiment such as drum rotation speed, and collecting data, using an example from our own research. This worked example can serve as a guide to researchers wishing to conduct their own optomotor experiments.

Generating a hypothesized acuity for the species of interest

We wanted to measure acuity in a species of swordtail fish (Xiphophorus spp.) where individuals are between 3 and 16cm in total length. In the literature, we found a published regression between body size (total length) and acuity in ray-finned fish (Caves, Sutton, & Johnsen, 2017). Using this regression, we predicted that acuity in the smallest individuals of our species would be approximately 3 cycles per degree (cpd), and in the largest individuals could be as high as 5 cpd. The published regression was, however, based on anatomical estimates of acuity (i.e. those produced using the density of the photoreceptors or retinal ganglion cells), which can give estimates of acuity that are higher than are found with behavioural methods, such as optomotor. Thus, to account for this, and to ensure that we tested a wide range of spatial frequencies likely to encompass the real acuity value, we created stimuli of spatial frequencies from 1 to 6 cpd.
We chose an experimental arena that was 300mm wide and which we could fill with water to a height of 100mm. This allowed sufficient water volume for the needs of our fish species. However, to keep the fish near the centre of the arena whilst still allowing it to move relatively freely, we also used a second cylinder (100mm diameter) to constrain the animal. Based on this setup, we required an optomotor apparatus capable of supporting an experimental arena that was 300mm in diameter (and thus stimuli that are slightly larger in diameter than that, to allow them to rotate freely around the arena) and weighed approximately 7kg.
Based on our experimental needs, the 3D printing templates provided here are for an apparatus that supports an arena 300mm in diameter and stimuli 340mm in diameter. However, these parts can support any diameter of arena and stimuli smaller than 340mm. For example, if performing optomotor experiments on a small animal (for example an insect), it may be prudent to choose a smaller arena in which to confine the animal. No changes to the design provided are necessary to scale down, and the stimulus calculation spreadsheet (see supplemental material) can calculate stripe widths for any drum diameter. On the other hand, testing an animal that is larger or requires more space may require one to scale up our design. The size of the 3D-printed components can be altered and scaled up using open-source editing software such as Blender. The upper size limit for our design is one of weight, rather than diameter, as the central column can only support up to approximately 7kg; however, additional weight can be accommodated by printing the base bracket with thicker walls, or in solid plastic.

Constructing and programming the optomotor apparatus

We designed the optomotor apparatus so that it could be constructed affordably from 3D printed parts, readily available components, and with only basic tools (i.e. hex-keys/allen-keys, a craft knife, and a computer for programming the microcontroller). A full list of parts and their approximate cost is shown in Table 1, and a full demonstration of the assembly process is shown in the supplementary online video (above). Here we provide a brief overview of the assembly process, although we recommend viewing the video for a more detailed guide. Parts can be downloaded here.

3D printing of components

The 3D printed parts can be constructed using a standard “fused filament fabrication” (also known as “fused deposition modelling”) printer. The 3D parts are available as individual .stl models for use with the printer’s slicing software (here). Any standard plastic is likely to function adequately, such as ABS, PLA or PETg. We used ABS due it its strength and flexibility, with 0.2mm layer depth and 15% infill for all models. An advantage of ABS is the ability to use acetone vapour baths to partially dissolve the surface and improve layer lamination strength (which makes the parts stronger, particularly under pressure near bolts).

Apparatus Assembly

See Figure 1 for schematics and photographs of the assembled system. The bottom bracket and motor housing attach to two aluminium profile sections (commonly sold as 2020 t-slot profile), holding them 100mm apart. T-slot nuts and 10mm M5 bolts are used to secure these parts to the aluminium profile. A 100mm M10 bolt fits through the bottom bracket, holding the sealed bearing unit in place, and above this sits the top bracket, secured with an M10 lock-nut. The bearing is sandwiched between a pulley/sprocket with 80 teeth, and the drum’s arm hub (connected around the bearing unit using M3 nuts and 20mm bolts). Four arms attach to the hub, creating spokes of the drum wheel (using M3 nuts and 10mm bolts), and four arc sections create the outside of the wheel (using M3 nuts and 20mm bolts). The pulley is designed to fit a 6mm GT2 belt, connecting to the stepper motor with a 20-tooth pulley. Sliding the bottom bracket or motor housing parts away from each other allows for belt tension adjustment. The motor and 20-tooth pulley can also be moved up and down to match the height of the 80-tooth pulley on the drum.

Arena Assembly

For the size of drum we required (300mm diameter), the base of the drum was a circle of extruded clear acrylic (PMMA), 10mm thick and 320mm in diameter. (Note that a waterproof material such as acrylic is required to construct an arena for an aquatic animal. In the supplemental video, we also provide an example of how to construct an arena for a terrestrial animal, in which case the material need not be waterproof and can be constructed of a material like foamed PVC of 5mm thickness, which can be cut with a craft knife.) The base must be a homogeneous, uniform colour with no visual rotational cues. To finish creating the arena, we acquired a 300mm long by 300mm diameter section of transparent acrylic tube, and glued this to the base using aquarium silicone. In the centre of this we positioned a second, internal tank made from 100mm diameter acrylic tube. This was secured to the bottom of the tank using a 3D printed part that allowed free water flow between the large external and small internal tank.

Electronics & Wiring

An optomotor drum requires smooth, slow, and precise rotation. We therefore used a stepper motor that offers control, and a combination of gearing and micro-stepping to achieve consistent fluid rotation at speeds lower than 1 revolution per minute (rpm). Micro-stepping is the use of graded opposing currents to hold the motor at intermediate angles between steps. The stepper motor is powered and controlled by a driver board, which is in turn given instructions by an Arduino microcontroller. The Arduino in turn receives user commands via an infrared remote control. Table 1 lists the parts we used, Figure 2 shows the wiring diagram, and the supplementary online video guide gives further specific construction instructions. There are many alternative Arduino boards or other microcontrollers which will be capable of running the drum. For example, a Raspberry Pi could be used to integrate your drum with functions provided by a full operating system, such as using on-the-fly video tracking (e.g. OpenCV), or a rotary encoder to control drum rotation. An additional alternative is to have the Arduino communicate with a standard computer via the USB port (using the serial connection). The electronics can be assembled using a solderless prototyping breadboard, meaning that no soldering is required. However, for long-term use (particularly in humid/damp environments) we recommend soldering all the contacts to make the system more robust. You may also wish to create a lid for the electronics housing in some environments.
The Arduino must be programmed by connecting it to a computer with a USB cable and using the free Arduino IDE software (supported on Windows, Mac or Linux). The code we used for controlling the Arduino is supplied along with the other files here, and demonstrates a range of functions. Our code requires two libraries be installed; “AccelStepper” and “Irremote”, both are freely available to download. The libraries should be copied across to your Arduino IDE “Libraries” directory on your computer. See the video guide for detailed instructions on the flashing process, and how to modify the code if your infrared remote is different to the one we used.

Description	Product code/description	Quantity	Net cost
Arduino + USB cable	UNO R3**	1	£10.00
Stepper motor	NEMA 17 Stepper motor, 17HS4401, 200 steps/revolution, 1.7A 40Ncm, 5mm shaft diameter	1	£11.00
Stepper motor driver	DRV8825	1	£3.00
Infrared receiver	VS1838B**	1	£0.50
Infrared remote	(Elegoo used, but many types will be compatible)**	1	£1.00
Aluminium Profile	2020 T-slot 20×20 approx. 600mm	2	£10.00
Sheet material	e.g. expanded PVC or painted plywood. Approx. 5mm thickness, roughly 400x800mm	1	£7.00
Diffuser	PTFE 320mm diameter, 0.5mm thick	1	£6.00
Drum walls	Card/paper with laser-printed stripes	4	£0.20
Light source	Focussed LED lamp (non-flickering)	1	£10.00
Camera	Basic 1080p “action camera”	1	£30.00
Ball bearing	Inside diam: 10mm, outside diam: 30mm depth:9mm	1	£2.50
M3 Bolts	M3 10mm bolt*	10	£0.50
M3 Bolts	M3 20mm bolt*	12	£0.50
M3 Nuts	M3 nuts*	22	£0.50
M5 Bolts	M5 10mm bolt*	8	£1.00
T-slot nuts	2020 M5 T-slot nut*	8	£7.00
M10 Bolt	M10 100mm bolt*	1	£1.00
M10 lock nut	M10 lock nut*	1	£0.25
M10 washers	M10 washers*	0-5	£0.25
Power supply	12v 2 amp (2000mA) power supply with 2.1mm connector	1	£10.00
Belt	GT2 6mm belt width, 420 teeth, 840mm	1	£5.00
Pulley/sprocket	GT2 6mm, 20 teeth, 5mm bore	1	£3.00
Prototyping board	Solderless breadboard, 170 points**	1	£1.50
Jumper leads	Various colours recommended**	17	£0.50
3D Printed parts
Arm hub	Holds arms in place	1	£0.30
Motor housing	Housing for stepper motor and electronics boards	1	£0.88
Arc section	Curved drum wheel sections	4	£1.88
Top bracket	Bracket for holding floor of drum	1	£0.60
Arm	Arms/spokes of drum wheel	4	£0.80
Pulley	Sprocket/pulley with 80 teeth and bearing hub	1	£0.23
Bottom bracket	Bracket securing base of drum	1	£0.73
* A2 stainless steel recommended for normal situations, or marine grade for use around salt water.
** These parts may be bought substantially cheaper as part of an Arduino kit

Stimulus construction

The drum is designed to accommodate stimuli made from four sheets of A4 paper, card, or waterproof-paper, although any flexible material could be used. Each sheet requires a small notch to be removed from the bottom corners so that it slots into the channel on each arc section of the wheel (see stimulus template file). Depending on the material you use to create the stimuli, they may be sturdy enough to stand alone. However, if they are not, they can be reinforced at the top using a ring of sturdy material such as cardboard.
To calculate how wide the stripes should be for stimuli of a given spatial frequency, we used the “Cycle width calculation supplement.xlsx” supplemental worksheet.

• 1. First, we filled in the diameter of the stimuli that we were going to use. Based on the size of our experimental arena (300mm diameter), we chose to use stimuli that were 340mm in diameter, so they had sufficient space to rotate freely around the arena. Therefore, we entered “340” into Box B7 of the spreadsheet.
• 2. Next, there are several ways the spreadsheet can calculate stripe widths for stimuli.
  • a. First, the spreadsheet can calculate the dimensions of a stimulus when given the desired spatial frequency in cycles per degree. To do so, enter the desired cycles per degree of a given stimulus in Box B21. This will produce the width of each stripe (Box C21) or stripe pair (i.e., cycle; Box D21) for that stimulus.
  • b. Second, the spreadsheet can calculate stripe widths for a large number of spatial frequencies. After entering the diameter of your stimuli in Box B7, the yellow table beginning in Line 31 will automatically populate with stimulus dimensions for a large number of spatial frequencies.
  • c. Third, the spreadsheet can calculate the spatial frequency (in cpd) of a stimulus (Box D25), given the width of a single stripe (Box B25). This functionality could be used, for example, if someone has previously created optomotor stimuli that you would like to use, but for which you do not know the spatial frequency.
• 3. Because we wanted to create stimuli that ranged in spatial frequency from 1 to 6 cpd, we used the yellow table starting in Line 31 to determine the dimensions of a large number of stimuli. Using the table, we found that we should create stimuli of the following dimensions:

Spatial frequency (cpd)	Width of single stripe (mm)	Width of stripe pair (cycle; mm)
1	1.49	2.98
2	0.75	1.49
3	0.5	0.99
4	0.37	0.75
5	0.3	0.6
6	0.25	0.5

• 4. We then created stimuli that comprised vertical black and white stripes of the thicknesses noted in the table above. These should be created as vector graphics (using a vector graphics software such as Adobe Illustrator or the open-source alternative Inkscape; see stimulus template file), and printed with vector rendering. We found, however, that when printing high-frequency stimuli, they exhibited distortions called Moiré patterns (where lines differ in relative size, angle or spacing). These distortions can arise due to the native resolution of the printer. For example, with the stimulus widths in the above table, the highest-frequency stimulus (6 cpd) exhibits Moiré patterns, where line thickness and spacing is uneven (Figure 3):

Therefore, to print stimuli that do not exhibit distortion, we had to account for the native resolution of the printer in choosing the precise stripe widths for the stimuli. To do so, we used the green table in columns G-I in the “Cycle width calculation supplement.xlsx” file. In the blue Box (H13), we entered the native resolution of the printer in dots per inch, which we found in the printer’s technical specifications. The spreadsheet then calculates the size of a single dot (in mm) that the printer can produce. To avoid distortions in the stimuli at high spatial frequencies, we would need to choose stimulus widths that are whole number multiples of the size of a single dot, which the spreadsheet lists in column H, starting at line 17.

For example, the native resolution of our printer is 1200 dots per inch, so we entered “1200” into Box H13. The spreadsheet then calculated that each dot the printer produces is 0.021mm (Box H15). When we tried to print a stimulus at exactly 6cpd (with the width of a single stripe being 0.25mm), distortions in the printed copy arose because 0.25 is not a whole number multiple of 0.021. Using the green table in the spreadsheet, however, we could see that a stimulus of 6.46 cpd can be made using stripe widths that are 0.231mm each. Thus, we could not create stimuli with this printer that are exactly 6cpd, but instead chose a value that was close to 6cpd and which could be printed without any distortions. Note that Moiré distortions tend to only be problematic at high spatial frequencies.

Following the next round of printing, we found no distortions in our stimuli. Thus, the final set of stimuli we printed included 1, 2, 3, 4, 5, and 6.46cpd. We then fashioned the printed stimulus sheets into cylindrical stimuli that could remain upright when placed around the test arena. We also created a “control” stimulus that was made out of plain grey cardstock.

Illumination

It is essential that the drum is well illuminated, and that the illumination from above is entirely consistent, with no shadows or shapes visible or cast onto the roof of the drum which might act as visual cues to the animal inside. We therefore recommend creating a diffuser sheet on top of the drum and using consistent illumination from above. We used a 0.5mm thick sheet of PTFE, cut to a 360mm diameter and attached to a 360mm ring of material such as foamed PVC or cardboard, to keep it flat and sturdy. However, other diffuse materials could be used, such as tissue paper, as long as they are suitably homogeneous in their light transmission. In the centre of the diffuse sheet there must be a circular hole for the video camera to see though.
The drum must be well illuminated, and ideally the light levels should be measured prior to the start of trials (e.g. with a light meter or spectroradiometer) and reported alongside any published results. One of the main requirements of the light source is that it does not flash at a speed that could interfere with the motion of the stripes on the stimuli. LEDs powered via low-voltage circuits are likely to be suitable, as are arc lamps, metal halide lamps, etc. Avoid the use of LEDs powered directly from mains electricity (these are often multiple LEDs wired together in series), because they will flash at the mains AC frequency (50Hz), which is slow enough to potentially cause considerable motion confusion for moving stripes. The presence of flashing can be tested with a high speed camera, capable of 120 frames per second or higher (this is available on many smart phones). Any flashing effects should be readily apparent in the high-speed video footage (see video guide for a demonstration). The light source must cast a uniform illumination onto the diffuser sheet from above, ensuring that any brightness gradients are uniform from the centre to the edges of the circular diffuser sheet.

Recording optomotor trials

The animal’s optomotor response to the drum rotation was recorded using a video camera. This: i) limits experimenter bias (particularly if the experimenter cannot see the animal’s reaction to the drum rotation), ii) acts as a valuable resource for objectively quantifying the animal’s response, and iii) creates a record for archiving and data transparency. The camera itself must be positioned in the centre of the diffuser sheet, however it would create a shadow of its outline. We therefore recommend making an opaque circular disk with a central hole just large enough for the lens to see through, and an external diameter large enough to completely block the camera’s shadow from reaching the diffuser. The animal looking up from the base of the drum will therefore only be able to see a circular shadow in the centre of the roof, with a circular lens looking through, offering no rotational movement cues.

Preliminary experiments: choosing experimental parameters

From the literature and Table 2 provided here, we found published drum rotation speeds in fish similar in size to our species of interest ranged from 1 to 15 rpm. However, preliminary experiments with our fish showed that they exhibit a robust optomotor response to a stimulus of 1cpd (our lowest-frequency stimulus) when presented at 1 rotation per minute. Thus, based on the recommendation that one should select the lowest rotation speed that elicits a reliable response, we chose to rotate the drum at 1rpm.
Preliminary experiments at 1 rpm and using various stimuli showed that individuals often took up to 30 seconds to respond to stimulus rotation; therefore, we programmed the apparatus to rotate for 45 seconds in each direction, which gave the fish sufficient time to respond to the stimulus after accounting for response latency. Additionally, these preliminary experiments allowed us to determine which optomotor response behaviours are robust and reliable in this species. For example, we found that individuals generally exhibit robust tracking responses in one direction, but weren’t very adept at changing direction and also generally continued to rotate for approximately 3-5 seconds after the stimulus stopped rotating.
To account for this, we decided to present each stimulus a total of four times: two clockwise rotations and two counter-clockwise rotations, and to define a positive optomotor response as at least one-half rotation or pivot of the whole body in the direction of drum rotation in at least two of the four trials for a given stimulus.

Collecting data

For each fish, we used the function Sample in the open-source R platform (R Development Core Team, 2019) to generate a random order in which to present the stimuli. This ensured that stimuli were not consistently presented from thicker to thinner stripes or vice versa. All trials were video recorded.

Data Analysis

Here, as a representative example, we display data for one fish from our experiment and discuss data analysis.
First, two viewers annotated all of the videos. By using a paper screen with a “viewing porthole” that showed only the fish, but not the rotating stimuli, we ensured that the viewers were blind to both the direction of stimulus rotation as well as to the stimulus identity. Based on Brokovich et al. (2010), Dobberfuhl, Ullmann, & Shumway (2005), and our own preliminary annotation of videos, we defined a positive response as at least one-half rotation by swimming or pivoting in the direction of drum rotation and at roughly the speed of stimulus rotation. Each fish was given four trials with each stimulus, and we considered that a fish had responded to that stimulus if it displayed positive responses in at least three out of four trials. Therefore, when annotating the videos, viewers noted whether or not the fish achieved at least ½ rotation in either the clockwise or counter-clockwise direction (see table 3 below). See Supplemental Video 2 for an example of a positive response at 0.4 cycles per degree, and Supplemental Video 3 for an example of a negative response at 4 cycles per degree. After annotating all of the videos, the annotations from the two viewers were compared; agreement between the two viewers was 90%, so the annotations from the viewer with more experience viewing optomotor footage was used for analysis.

Trial Number	Rotation Direction	Stimulus Frequency	Positive Response?
1	CL	0.2	1
2	CO	0.2	1
3	CL	0.2	1
4	CO	0.2	1
1	CL	0.4	1
2	CO	0.4	1
3	CL	0.4	1
4	CO	0.4	1
1	CL	0.6	1
2	CO	0.6	1
3	CL	0.6	1
4	CO	0.6	0
1	CL	0.8	1
2	CO	0.8	1
3	CL	0.8	0
4	CO	0.8	1
1	CL	1	0
2	CO	1	1
3	CL	1	1
4	CO	1	1
1	CL	2	1
2	CO	2	1
3	CL	2	0
4	CO	2	1
1	CL	3	1
2	CO	3	1
3	CL	3	0
4	CO	3	1
1	CL	4	0
2	CO	4	1
3	CL	4	0
4	CO	4	1
1	CL	5	0
2	CO	5	0
3	CL	5	1
4	CO	5	0
1	CL	6	0
2	CO	6	0
3	CL	6	0
4	CO	6	0
1	CL	control	0
2	CO	control	0
3	CL	control	0
4	CO	control	0

We then plotted the number of trials in which a fish performed at least a half-turn for each stimulus (Figure 4). From the figure, it can be seen that the criteria for a positive response were met at 0.2, 0.4, 0.6, 0.8, 1, 2, and 3 cycles per degree. However, the responses fell below the criteria for a positive response at 4, 5, and 6 cycles per degree. In no trials did the fish display at least a half-rotation to the control stimulus. Therefore, based on these data, we conclude that the acuity of this fish is between 3 and 4 cycles per degree (0.25 degrees minimum resolvable angle).

Although similar data are not available for this species from any other studies, this value does align well with predictions based on the size of the fish, which has been previously shown to be a very strong predictor of acuity across ray-finned fish (Caves et al., 2017). In particular, based on the fish’s body size (3.27cm), we would predict that its acuity is 3.48cpd, which is consistent with the values we recorded using the optomotor apparatus.

Class	Genus	Species	Common name	Drum Speed (rpm)	Stimulus Diameter (cm)	Internal Cylinder Diameter (cm)	Animal restrained?	Light Adaptation Time (min)	Source
Actinopterygii	Danio	rerio	Zebrafish	02/05/20	39		Yes		(Tappeiner et al., 2012)
Actinopterygii	Dascyllus	marginatus	Damselfish	2.25	41	20	No	20	(Brokovich et al., 2010)
Actinopterygii	Enantiopus	melanogenys	Cichlid	4	28	24.5	No	10	(Dobberfuhl et al., 2005)
Actinopterygii	Forsterygion	varium	Variable triplefin (larvae)	0-8	6 or 12	2	No		(Pankhurst, Pankhurst, & Montgomery, 1993)
Actinopterygii	Oncorhynchus	mykiss	Rainbow trout (larvae)	8	6 or 12	2	No	1	(Rahmann, Jeserich, & Zeutzius, 1979)
Actinopterygii	Oryzias	latipes	Medaka	4	48	0.6-9	No		(Carvalho, Noltie, & Tillitt, 2002)
Actinopterygii	Pleuronectes	platessa	Plaice (larvae)	2-15	9.5 or 15.5	2	No		(Neave, 1984)
Actinopterygii	Xenotilapia	flavipinnis	Yellow sand cichlid	4	28	24.5	No	10	(Dobberfuhl et al., 2005)
Actinopterygii	Xenotilapia	leptura	Cichlid	4	28	24.5	No	10	(Dobberfuhl et al., 2005)
Actinopterygii	Xenotilapia	spilopterus	Cichlid	4	28	24.5	No	10	(Dobberfuhl et al., 2005)
Amphibia	Rana	temporaria	Common frog	6	44				(Birukow, 1937)
Amphibia	Triturus	cristatus	Crested newt	1	74, 58, 48, or 20		No		(Manteuffel & Himstedt, 1978)
Aves	Calypte	anna	Anna’s hummingbird	1	100		Yes	2	(Goller et al., 2019)
Aves	Gallus	gallus	Domestic Chicken (chick)	1	50		Yes		(Schmid & Wildsoet, 1998)
Branchiopoda	Daphnia	pulex	Water flea	1.22-4	24	4	No	02/04/20	(Hathaway & Dudycha, 2018)
Cephalopoda	Nautilus	pompilius	Nautilus	4	50	25	No		(Muntz & Raj, 1984)
Cephalopoda	Octopus	vulgaris	Common octopus	1	30 or 60	7.5	No		(Packard, 1969)
Cephalopoda	Sepia	officinalis	Common cuttlefish	3	152	23	No	1	(Groeger, Cotton, & Williamson, 2005)
Insecta	Apis	mellifera	Honeybee	5	15.5		Yes		(Moore, Penikas, & Rankin, 1981)
Insecta	Hypothenemus	hampei	Coffee Berry Borer	13	31		No		(Vega et al., 2014)
Insecta	Xenos	peckii	Twisted-wing insect		16		Yes		(Maksimovic, Layne, & Buschbeck, 2007)
Malacostraca	Alpheus	heterochaelis	Snapping shrimp		30	7.5	No		(Kingston, Lucia, Havens, Cronin, & Speiser, 2019)
Malacostraca	Ancylomenes	pedersoni	Pederson shrimp	6	56	8	No	15	(Caves, Frank, & Johnsen, 2016)
Malacostraca	Callinectes	sapidus	Blue crab	2	56	30	No	15	(Baldwin & Johnsen, 2011)
Malacostraca	Lysmata	amboinensis	Scarlet cleaner shrimp	6	56	8	No	15	(Caves et al., 2016)
Malacostraca	Pachygrapsus	crassipes	Striped shore crab	0.5	48		Yes		(Wiersma & Hirsh, 1974)
Malacostraca	Panulirus	interruptus	rock lobster	0.5	48		Yes		(Wiersma & Hirsh, 1974)
Malacostraca	Procambarus	clarkii	Crayfish	0.5	48		Yes		(Wiersma & Hirsh, 1974)
Malacostraca	Urocaridella	antonbrunii	Clear cleaner shrimp	6	56	8	No	15	(Caves et al., 2016)
Mammalia	Anoura	geoffroyi	Geoffroy’s tailless bat	6	40	12	No		(Suthers, 1966)
Mammalia	Antrozous	pallidus	Pallid bat	01/05/20	38	12	No	30	(Bell & Fenton, 1986)
Mammalia	Artibeus	jamaicensis	Jamaican fruit bat	6	40	12	No		(Suthers, 1966)
Mammalia	Carollia	perspicillata	Seba’s short-tailed bat	6	40	12	No		(Suthers, 1966)
Mammalia	Desmodus	rotundus	Common vampire bat	6	40	12	No		(Suthers, 1966)
Mammalia	Diaemus	youngi	White-winged vampire bat	6	40	12	No		(Suthers, 1966)
Mammalia	Eptesicus	fuscus	Big brown bat	01/05/20	38	12	No	30	(Bell & Fenton, 1986)
Mammalia	Macrotus	californicus	California leaf-nosed bat	01/05/20	38	12	No	30	(Bell & Fenton, 1986)
Mammalia	Monodelphis	domestica	Short-tailed opossum	2.4	56	15	No		(Dooley, Nguyen, Seelke, & Krubitzer, 2012)
Mammalia	Mus	musculus (lab)	Lab mouse	2	29	11.5 (platform)	No	10 (scotopic1); 5 (photopic2)	(Abdeljalil et al., 2005)
Mammalia	Mus	musculus (lab)	lab mouse	2	30	8	No	0.5	(Thaung, Arnold, Jackson, & Coffey, 2002).
Mammalia	Myotis	lucifugus	Little brown bat	6	40	12	No		(Suthers, 1966)
Mammalia	Oryctolagus	cuniculus	Rabbit	01/03/20	63.5	50 (platform)	Yes		(Hahnenberger, 1977)
Mammalia	Peromyscus	leucopus	White-footed mouse	2.5-6	20, 40, or 80	12.5 (platform)	Yes	2	(Vestal, 1973)
Mammalia	Peromyscus	maniculatus	Deer mouse	2.5-6	20, 40, or 80	12.5 (platform)	Yes	2	(Vestal, 1973)
Mammalia	Phyllostomus	hastatus	Greater spear-nosed bat	6	40	12	No		(Suthers, 1966)
Mammalia	Rattus	norvegicus	Rat	2	58	22 (platform)	No	0.5	(Cowey & Franzini, 1979)
Mammalia	Rattus	norvegicus	Rat	2	48		No		(Douglas et al., 2005)
Mammalia	Saccopteryx	bilineata	Greater sac-winged bat	6	40	12	No		(Suthers, 1966)
Mammalia	Saccopteryx	leptura	Lesser sac-winged bat	6	40	12	No		(Suthers, 1966)
Reptilia	Chamaeleo	chamaeleon	Common chameleon	1	136 or 93	18	Yes for OKR3; no for OMR4	0.5	(Lev-Ari, Katz, Lustig, & Katzir, 2017)
Reptilia	Chelonia	mydas	Green turtle (hatchlings)	10	38	19	No	5	(Ireland, 1979)
Reptilia	Pseudemys	scripta	red-eared turtle	20	30		Yes	10	(Hayes & Ireland, 1972)
Reptilia	Ptyodactylus	hasselquistii	Fan-footed gecko	2-11.5	50	20	No	15	(Frankenberg, 1981)
Reptilia	Sphenodon	punctatus	Tuatara	4	102.5	50.8	No	5	(Ireland & Gans, 1977)

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