# Photographing Butterflies in the Lab

[EcoEvoDevoLab](https://hackmd.io/@EcoEvoDevoLab/AngeliniLab)

Updated 8 March 2022


Butterflies caught in the wild or raised in the lab can be studied for variations in wing shape and pattern. First, it is necessary to photograph them under consistent conditions.

## Relaxing dry butterfly specimens

Butterflies collected dead in the field or from lab stocks typically dry in orientations that don't allow good photography. Thankfully, they can be "relaxed" by exposure to high humidity and repositioned before imaging. There is a danger that the high humidity will encourage mold growth. So it is best to do this procedure no longer than over-night. Ideally, you can practice with low-value specimens, before trying to relax rare specimens.

Find a suitable chamber large enough to tightly enclose butterfly specimens and liquid water, without the two touching. I have had success with an old plastic croissant tray sold at Walmart.

- Place a Petri dish filled with wet paper towel in the humidifying chamber.
- Place one or more butterflies in a second, dry Petri dish.
- Close and seal the lid of the chamber.
- Incubate for up to 24 hours or until the specimens can be repositioned without breaking.


## Spreading butterflies

To get photographic data that includes all of the forewings and the hindwings, butterflies should be pinned with their wings spread out. For instructions on this process, first watch this 5-minute [video](https://www.youtube.com/watch?v=f-QIfygvhhM).


A few modifications that are especially helpful if you are spreading several butterflies at once:

- Start at the top of the spreading board, far from you. If you spread multiple butterflies at once, place later specimens closer towards you. This avoids the possibility that you will accidentally crush a specimen while leaning over it.
- You can use small strips of vellum (translucent paper) rather than a big sheet, as shown in the video. The strips should be long enough that they can secure the front and back wings on one side. For most *V. cardui*, that's a peice about 7 cm by 2 cm.
- Don't worry about the antennae! -- If you want to be fancy, go ahead and position them. But our lab doesn't use data from antennae in any research projects (right now).
- Rodents will eat pinned insects! Right now, we don't have a great place to allow them to dry. Right out on the bench is probably okay. Since there's plenty of day time traffic to scare off mice.
- Drying time will vary with the ambient humidity. A week is ideal, but about 3 days may be sufficient.

## Labeling specimens

Image data are not useful without metadata explaining where specimens come from.

### Field-caught specimens

All insects caught in the field should immediately be given a specimen ID number. This number is based on the initials of the person making the collection and the date of capture. It begins with 3 initials of the collector's name (e.g. DRA), two digits for the year, two digits for the month, two digits for the day of the month, a dash, then 3 digits which are arbitrary, but meant to simply count up as a person collects multiple specimens on the same day. This number should be written on the collecting vial and in a field notebook. The field notebook should record any relevant metadata, such as the location, time of day, weather, etc. and a preliminary taxonomic identification. The number should then accompany that specimen (and any samples derived from it, like DNA) in perpetuity.


Once a specimen enters an insect collection and it is pinned, some critical metadata should be placed on a paper label that can also be held on the pin. Labels can be hand-written, but ideally they should be printed from a computer. Size 4 font works for this purpose! The first label should carry the specimen ID number. The second label carries "locality" information: the country, region (state or province), county, city or town and street address or local name. Latitude and longitude (GPS coordinates) should be included to 4 decimal places. Then the date of collection and the collector's name should appear.

Eventually, someone should identify the specimen! This isn't always possible, and many collections have unidentified specimens. These should nevertheless have locality tags. If a taxonomic identification is made, a species label can be added to the stack of paper pinned below the insect. ID labels should include the species name and common name (if there is one), the order and family and the name of the person making the determination.

### Lab-reared specimens

In our lab, specimens raised in captivity still get an ID number. The true bugs have a separate convention we've developed. But for butterflies, we will adapt the field entomology system: Use "LAB" in pace of the collector's initials and the date on which the tags are prepared. For the final 3 digits, it will helpful to keep count over multiple days, so that all of the specimens from one experiment have non-repeating, consecutive numbers.

#### *Vanessa cardui*

For painted lady butterflies, it will be helpful to give them specimen ID numbers after they have been spread and dried, before being photographed. At this time, also determine the sex of the specimen, and record that information on the tag with a male ♂ or female ♀ symbol. Males are typically smaller than females, although that is not a [reliable way](https://twitter.com/Aphanotus/status/1328834844269359107) to distinguish them. Check the ends of the front legs under a microscope or using a magnifying glass: males have very hairy feet, while females have 3 rows of claws.


## Photographing butterfly specimens

Once you have pinned, spread, labeled butterflies, you're ready to take some pictures!

:::info
Check out the [tutorial on field photography](https://hackmd.io/@aphanotus/macrophotography?type=view), which explains [shutter speed](https://hackmd.io/@aphanotus/macrophotography#Shutter-speed), [aperture](https://hackmd.io/@aphanotus/macrophotography#Aperture) and [ISO](https://hackmd.io/@aphanotus/macrophotography#ISO), and [how to use them](https://hackmd.io/@aphanotus/macrophotography#Aperture-ISO-shutter-speed).
:::

To ensure consistent photographic conditions in the lab, we will use a fixed camera set-up. (Butterflies are too large to use a microscope-mounted camera.) It is best not to adjust the tripod or remove the camera or flash.


- **Turn on the camera**. The on/off switch is on the upper left of the camera back.
- Check that the camera is in **manual mode**. The upper right thumb wheel should be set to "M". And if you press the "Q" button in the upper left, the screen should look like it does in this example, with a large blue "M".
- Check the shutter speed, aperture and ISO **settings**. Those shown in the picture below should be good for most specimens.
- **Turn on the flash** by holding down the on/off button for 3 seconds. If the flash has an adjustment for distance


- **Position the specimen** below the camera on a plain styrofoam board. Look through the eyepiece and adjust the specimen so that:
1. the specimen is **centered**, and all anatomical parts of interest are in the view
2. the specimen **ID label** is visible
3. the metric **ruler** and color standard are visible


- **Focus** the lens by gently pressing down the shutter release (the smooth button on the top/front right of the camera).
- When you're ready to take a photograph, **press the shutter release button** all the way down. You will hear the internal mechanism of the camera, and the image will display on the camera's screen.
- Confirm that the image is focused and [properly exposed](https://photographyhero.com/wp-content/uploads/2017/05/images-exposure-compensation-range.jpg). If necessary, refocus or [adjust the settings](https://hackmd.io/@aphanotus/macrophotography#Camera-Basics), and take the picture again.
- After you have a dorsal image, **flip the specimen over** (push the head of the pin into the styrofoam board), and image the ventral wing surface.

:::info
It's also possible to use a [remote trigger](https://www.youtube.com/watch?v=yBsNgyDwhJA) on the camera. This will reduce vibration, which is especially helpful if the manual settings use a low ISO and slow shuttle speed.
:::

### Moving image data off the camera

- After a session of photography, **turn the camera and flash off**.
- If you have a laptop, **plug the USB cable running from the camera into your computer**.
- **Turn on the camera**. It will appear as an external device.
- **Log on** to Colby's file-share system or [NSCC](https://www.colby.edu/arc/hardwaresystems/computer-clusters/nscc/), Colby's high-performance computing cluster.

:::info
If you don't have access to NSCC, [email Dave](mailto:[e-mail])!
:::

- If necessary create a new folder under `nscc:/research/drangeli/`. Be sure to name the folder something meaningful for your project.
- **Copy image files from the camera to the research folder**. You may also choose to keep the files on your laptop.
- Confirm that the files are at their destination, then **delete the images from the camera**.

:::info
Ideally the name of each image file includes the specimen ID and other important metadata. However, renaming many files can be tedious. If it's obvious from the folder name what project the images are for, and the specimen ID and sex are visible in the label in the image, then don't worry about renaming the files.
:::

## What's next?

So what happens to all these images?

Images can be used qualitatively to document naturally occuring or experimentally induced phenotypes. Various methods exist for morphometric analysis (the quantification and comparison of shape) and colorimetry (quantification and comparison of color). These and other topics are explored in [BI377 Morphometry](https://github.com/aphanotus/openEd/tree/main/BI377.22F.morphometry#bi377--morphometry-the-study-of-form).

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[EcoEvoDevo Lab](https://hackmd.io/@EcoEvoDevoLab/AngeliniLab)