Name: automated charting of the visual space of housefly compound eyes
Uploaded: 2022-03-31T14:20:00
Description: Watch this Scientific Journal Video about Automated Charting of the Visual Space of Housefly Compound Eyes at JoVE.com

This study was financially supported by the Air Force Office of Scientific Research/European Office of Aerospace Research and Development AFOSR/EOARD (grant FA9550-15-1-0068, to D.G.S.). We thank Dr. Primo&#382; Pirih for many helpful discussions and Kehan Satu, Hein Leertouwer, and Oscar Rinc&#243;n Carde&#241;o for assistance.

The protocol is in accordance with the University&#39;s insect care guidelines.
1. Preparation of a housefly,&#160;&#160;Musca domestica
<ol>
	<li>Collect the fly from the laboratory-reared population. Place the fly in the brass holder (Figure 1).
	<ol>
		<li>Cut 6 mm from the upper part of the restraining tube (see Table of Materials). The new upper part of the tube has an external diameter of 4 mm and an internal diam...

<ol>
	<li>Land, M. F., Nilsson, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Animal Eyes. , (2012).</li><li>Cronin, T. W., Johnsen, S., Marshall, N. J., Warrant, E. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Visual Ecology. , (2014).</li><li>Horridge, G. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+separation+of+visual+axes+in+apposition+compound+eyes.">The separation of visual axes in apposition compound eyes.</a> Philosophical Transactions of the Royal Society of London. Series B. 285 (1003), 1-59 (1978).</li><li>Land, M. 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The spatial distribution of the visual axes of housefly eyes can be charted using the pseudopupil phenomenon of compound eyes and the reflection changes caused by the light-dependent pupil mechanism. Therefore, an investigated fly is mounted in a goniometric system, which allows inspection of the local facet pattern with a microscope setup equipped with a digital camera, all under computer control. Image analysis yields eye maps. An essential difficulty encountered is that without careful positioning of the eye at the be...

The compactness of insect visual systems and the agility of their owners, demonstrating highly developed visual information processing, have intrigued people from both scientific and non-scientific backgrounds. Insect compound eyes have been recognized as powerful optical devices enabling acute and versatile visual capacities1,2. Flies, for instance, are well-known for their fast responses to moving objects, and bees are famous for possessing color vision and polarization vision2.
The compound eyes of arthropods consist of numerous anatomically similar units, the ommatidia, each of which is capped by a facet lens. In Diptera (flies), the assembly of facet lenses, known collectively as the cornea, often approximates a hemisphere. Each ommatidium samples incident light from a small solid angle with half-width on the order of 1&#730;. The ommatidia of the two eyes together sample approximately the full solid angle, but the visual axes of the ommatidia are not evenly distributed. Certain eye areas have a high density of visual axes, which creates a region of high spatial acuity, colloquially called a fovea. The remaining part of the eye then has a coarser spatial resolution3,4,5,6,7,8,9.
A quantitative analysis of the optical organization of the compound eyes is crucial for detailed studies of the neural processing of visual information. Studies of the neural networks of an insect&#39;s brain10 often require knowledge of the spatial distribution of the ommatidial axes. Furthermore, compound eyes have inspired several technical innovations. Many initiatives to produce bio-inspired artificial eyes have been built on existing quantitative studies of real compound eyes11,12,13. For instance, a semiconductor-based sensor with high-spatial resolution was designed based on the model of insect compound eyes11,14,15,16,17. However, the devices developed so far have not implemented the actual characteristics of existing insect eyes. Accurate representations of insect compound eyes and their spatial organization will require detailed and reliable data from natural eyes, which is not extensively available.
The main reason for the paucity of data is the extreme tediousness of the available procedures for charting the eyes&#39; spatial characteristics. This has motivated attempts to establish a more automated eye mapping procedure. In a first attempt at automated analyses of insect compound eyes, Douglass and Wehling18 developed a scanning procedure for mapping facet sizes in the cornea and demonstrated its feasibility for a few fly species. Here we extend their approach by developing methods for not only scanning the facets of the cornea but also assessing the visual axes of the ommatidia to which the facets belong. We present the case of housefly eyes to exemplify the procedures involved.
The experimental setup for scanning insect eyes is: partly optical, i.e., a microscope with camera and illumination optics; partly mechanical, i.e., a goniometer system for rotating the investigated insect; and partially computational, i.e., use of software drivers for the instruments and programs for executing measurements and analyses. The developed methods encompass a range of computational procedures, from capturing images, choosing camera channels, and setting image processing thresholds to recognizing individual facet locations via bright spots of light reflected from their convex surfaces. Fourier transform methods were crucial in the image analysis, both for detecting individual facets and for analyzing the facet patterns.
The paper is structured as follows. We first introduce the experimental setup and the pseudopupil phenomenon-the optical marker used to identify the visual axes of the photoreceptors in living eyes19,20,21. Subsequently, the algorithms used in the scanning procedure and image analysis are outlined.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Digital Camera</td>
			<td>PointGrey</td>
			<td>BFLY-U3-23S6C-C</td>
			<td>Acquision of amplified images and digital communication with PC</td>
		</tr>
		<tr>
			<td>High power star LED</td>
			<td>Velleman</td>
			<td>LH3WW</td>
			<td>Light source for observation and imaging the compound eye</td>
		</tr>
		<tr>
			<td>Holder for the investigated fly</td>
			<td>University of Groningen</td>
			<td></td>
			<td>Different designs were manufactured by the university workshop</td>
		</tr>
		<tr>
			<td>Linear motor</td>
			<td>ELERO</td>
			<td>ELERO Junior 1, version C</td>
			<td>Actuates the upper microscope up and down. (Load 300N, Stroke speed 15mm/s, nominal current 1.2A)</td>
		</tr>
		<tr>
			<td>Low temperature melting wax</td>
			<td>various</td>
			<td></td>
			<td>The low-temperature melting point wax serves to immobilize the fly and fix it to the holder</td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Zeiss</td>
			<td></td>
			<td>Any alternative microscope brand will do; the preferred objective is a 5x</td>
		</tr>
		<tr>
			<td>Motor and LED Controller</td>
			<td>University of Groningen</td>
			<td>Z-o1</td>
			<td>Designed and built by the University of Groningen and based on Arduino and Adafruit technologies.</td>
		</tr>
		<tr>
			<td>Motorized Stage</td>
			<td>Standa (Vilnius, Lithuania)</td>
			<td>8MT175-50XYZ-8MR191-28</td>
			<td>A 6 axis motorized stage modified to have 5 degrees of freedom.</td>
		</tr>
		<tr>
			<td>Optical components</td>
			<td>LINUS</td>
			<td></td>
			<td>Several diagrams and lenses forming an epi-illumination system (see Stavenga, Journal of Experimental Biology 205, 1077-1085, 2002)</td>
		</tr>
		<tr>
			<td>PC running MATLAB</td>
			<td>University of Groningen</td>
			<td></td>
			<td>The PC is able to process the images of the PointGrey camera, control the LED intensity, and send control commants to the motor cotrollers of the system</td>
		</tr>
		<tr>
			<td>Power Supply (36V, 3.34A)</td>
			<td>Standa (Vilnius, Lithuania)</td>
			<td>PUP120-17</td>
			<td>Dedicated power supply for the STANDA motor controllers</td>
		</tr>
		<tr>
			<td>Soldering iron</td>
			<td>various</td>
			<td></td>
			<td>Used for melting the wax</td>
		</tr>
		<tr>
			<td>Stepper and DC Motor Controller</td>
			<td>Standa (Vilnius, Lithuania)</td>
			<td>8SMC4-USB-B9-B9</td>
			<td>Dedicated controllers for the STANDA motorized stage capable of communicating with MATLAB</td>
		</tr>
		<tr>
			<td>Finntip-61</td>
			<td>Finnpipette Ky, Helsinki</td>
			<td>FINNTIP-61, 200-1000&#956;L</td>
			<td>PIPETTE TIPS FOR FINNPIPETTES, 400/BOX. It is used to restrain the fly</td>
		</tr>
		<tr>
			<td>Carving Pen Shaping/Thread Burning Tool</td>
			<td>Max Wax</td>
			<td></td>
			<td>The tip of the carving pen is designed to transfer wax to the head of fly</td>
		</tr>
		<tr>
			<td>MATLAB</td>
			<td>Mathworks, Natick, MA, USA</td>
			<td>main program plus Image Acquisition, Image Analysis, and Instrument Control toolboxes.</td>
			<td>Programming language used to implement the algorithms</td>
		</tr>
	</tbody>
</table>

automated charting of the visual space of housefly compound eyes

This paper describes the automatic measurement of the spatial organization of the visual axes of insect compound eyes, which consist of several thousands of visual units called ommatidia. Each ommatidium samples the optical information from a small solid angle, with an approximate Gaussian-distributed sensitivity (half-width on the order of 1&#730;) centered around a visual axis. Together, the ommatidia gather the visual information from a nearly panoramic field of view. The spatial distribution of the visual axes thus determines the eye's spatial resolution. Knowledge of the optical organization of a compound eye and its visual acuity is crucial for quantitative studies of neural processing of the visual information. Here we present an automated procedure for mapping a compound eye's visual axes, using an intrinsic, in vivo optical phenomenon, the pseudopupil, and the pupil mechanism of the photoreceptor cells. We outline the optomechanical setup for scanning insect eyes and use experimental results obtained from a housefly, Musca domestica, to illustrate the steps in the measurement procedure.

Animals and optical stimulation 
Experiments are performed on houseflies (Musca domestica) obtained from a culture maintained by the Department of Evolutionary Genetics at the University of Groningen. Before the measurements, a fly is immobilized by gluing it with a low-melting-point wax in a well-fitting tube. The fly is subsequently mounted on the stage of a motorized goniometer. The center of the two rotary stages coincides with the focal point of a microscopic setup24<...

Watch this Scientific Journal Video about Automated Charting of the Visual Space of Housefly Compound Eyes at JoVE.com

Automated Charting of the Visual Space of Housefly Compound Eyes

The protocol here describes the measurement of the spatial organization of the visual axes of housefly eyes, mapped by an automatic device, using the pseudopupil phenomenon and the pupil mechanism of the photoreceptor cells.

This paper describes the automatic measurement of the spatial organization of the visual axes of insect compound eyes, which consist of several thousands ...

The system and the protocol are designed to analyze the physio system of flies by mapping with minimal human intervention. With the system and the protocol, we can precisely know how the visual space of a fly size is organized. The system's advantages are the reproducibility and the speed of the mapping.The study of compound eyes is an important part of animal vision research and has inspired several technical innovations which have produced artificial eyes. We have set an example on how to build and test an automatic device for scanning compound eyes. Details of algorithm development that bring the parts together need particular attention.Start with collecting a fly from the laboratory-reared population. Prepare a restraining tube by cutting six millimeter from the upper part so that the tube has an external diameter of four millimeters and an internal diameter of 2.5 millimeters in the upper part. Place the fly inside the cut tube and seal the tube with cotton to prevent damaging the fly.Then push the fly such that the head protrudes from the tube and the body is restrained in the tube. Use beeswax to immobilize the head while the eyes remain uncovered. Once done, cut the tube to achieve a length of 10 millimeters.Then place the plastic tube containing the fly in the brass holder with one eye of the fly pointing upward and the holder resting on a tabletop. Adjust the orientation of the tube on the microscope as described in the manuscript to scan the whole eye within the range of the azimuth and elevation allowed by the setup. Set up the microscope by mounting an alignment pin on the azimuth rotation stage so that the X-Y position of the tip can be adjusted to coincide with the azimuth axis on the motorized stage.While viewing with the microscope equipped with a 5X objective, use the Z-axis joystick to focus on the tip. Next, align the X-Y adjustment of the azimuth axis with the microscope's optical axis and use the X and Y axis joysticks to ensure that the elevation and azimuth rotary axes are pre-aligned with the centered pin. Manipulate the azimuth and elevation joysticks to check whether the pin is centered with respect to both degrees of freedom.When well centered, the pin tip remains in the same position during azimuth and elevation rotations. Align and mount the fly with the elevation stage at zero degrees and holder on the azimuth stage. Then observe the fly's eye with the microscope.After turning the illumination LED on, adjust the horizontal position of the fly to align the center of the pseudopupil. Then change the vertical position of the pseudopupil by using the rotating screw of the holder so that the deep pseudopupil is brought into focus at the level of the elevation axis. Next, line the deep pseudopupil with respect to the azimuth and elevation axes by centering it in the field of view.When the setup is ready, switch the view to the digital camera mounted at the microscope and run the software initialization of the grace system, which includes initializing the motor controllers and the Arduino LED controller. To do so, open MATLAB version 2020a or higher version and run the MATLAB script. On the computer screen, confirm that the fly's pseudopupil at the center of the projected image.Use the Z-axis joystick to bring the focus to the level of the corneal pseudopupil. Once the focus is aligned, run the auto focusing algorithm to obtain a sharp image at the cornea level. Then return the focus to the deep pseudopupil level by adjusting the motorized Z-axis stage.Store the distance between the deep pseudopupil and corneal pseudopupil. Next, fine tune the pseudopupil centering with the auto centering algorithm followed by bringing the focus back to the corneal pseudopupil level. Rerun the auto focusing algorithm and zero the motorized stages at their current positions.While scanning the eye, run the scanning algorithm to sample the eye images along the trajectories in five degree steps while performing the auto centering and auto focusing algorithms. After the sampling, turn off the LED and motor controllers. Later, process the images by applying the image processing algorithms.In the study of optics of the fly eye, the image at the eye surface level shows the facet reflections and the pigment granule reflection in the activated state. The image taken at the level of the center of eye curvature illustrated the reflection of the arrangement of the photoreceptor cells in a trapezoidal pattern with their distal ends positioned at about the focal plane of the facet lenses. Two successive images were correlated to determine a shift in the translation of the facet pattern.An image taken during a scan across the eye is shown with the facet centroids. After an azimuthal rotation of five degrees, the subsequent image is illustrated here. The centroid procedure could not identify all the facets.A low local reflectance caused by minor surface irregularities, or specs of dust, resulted in erroneous centroids. The error was resolved by calculating a fast Fourier transform. The first ring of harmonics defines three orientations indicated by the blue, red, and green lines.The inverse transformation of the harmonics along the three orientations yielded the gray bands. The right eye of a Housefly was scanned from the frontal side to the lateral side in 24 steps. The image shows the assembly of the facets as a Voronoi diagram.At the start of the scanning, special attention should be given to the adjustment of the deep pseudopupil of the fly eye at the rotation center of the goniometric system. Here we apply epi-illumination microscopy. This method can be straightforwardly extended to fluorescence microscopy to study insects that do not have a reflecting pseudopupil.Quantitative knowledge of the distribution of the visual axes of an eye will allow understanding of how physio systems are optimized for certain tasks such as hunting, mating, or detecting predators.

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Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a ...

Trajectory Data Analyses for Pedestrian Space-time Activity Study

It is well recognized that human movement in the  spatial and temporal dimensions has direct influence on disease transmission1-3.  An infectious disease ...

Design and Analysis of Temperature Preference Behavior and its Circadian Rhythm in Drosophila

Design and Analysis of Temperature Preference Behavior and its Circadian Rhythm in Drosophila

The circadian clock regulates many aspects of life, including sleep, locomotor activity, and body temperature (BTR) rhythms1,2. We recently identified a ...

Capture Compound Mass Spectrometry - A Powerful Tool to Identify Novel c-di-GMP Effector Proteins

Considerable progress has been made during the last decade towards the identification and characterization of enzymes involved in the synthesis ...

Laser Capture Microdissection of Highly Pure Trabecular Meshwork from Mouse Eyes for Gene Expression Analysis

Laser capture microdissection (LCM) has allowed gene expression analysis of single cells and enriched cell populations in tissue sections. LCM is a great ...

Measuring the Confluence of iPSCs Using an Automated Imaging System

This study focuses on understanding how growing iPSCs on different ECM coating substrates can affect cell confluence. A protocol to assess iPSC confluence ...