This work was supported by grants from the U.S. Fish and Wildlife Service&#39;s Conservation Recovery Initiative (F17AP00467) and the Disney Conservation Fund. Additional support was provided by the Florida Museum of Natural History and the Department of Entomology and Nematology at the University of Florida.

1. Securing successful adult courtship and mating
<ol>
	<li>Following successful eclosion, release viable adult butterflies into a secure, walk-in, screened flight cage located in a temperature-controlled greenhouse (Supplementary Figure 2). 
	NOTE: Adults can be marked on the ventral surface of the wings with permanent ink markers if identification of specific individuals is desired for separation of genetic lines, stock origin, or for specific data collection related to organism longevity, behavior, etc.
	<ol>
		<li>While the exact cage dimensions can vary, ensure that there is ample space to accommodate adequate nectar plant material necessary to support the density of housed adult butterflies and provide room for a human to freely stand and pivot around.</li>
		<li>Beyond temperature regulation, ensure that the greenhouse is secure so that it can provide a second layer of containment along with protection from inclement weather (e.g., heavy rain, wind).</li>
	</ol>
	</li>
	<li>Elevate potted nectar plant material so that there is no more than 30 cm of space from the interior top of the cage to the highest blooming flowers (Supplementary Figure 2). This provides optimal access to the available nectar resources, offers ample adult perches, and minimizes extraneous flight activity.</li>
	<li>Place one potted host plant in the flight cage. This ensures that even if a mated pair is missed any resulting eggs laid can be collected.</li>
	<li>Provide consistent airflow. This enhances courtship activity and mating success. In a greenhouse setting, blowers and fixed mount circulation fans are best used to help enhance ventilation and air movement. Smaller portable ventilation, such as box or desk fans, can also be used.</li>
	<li>Maintain the internal greenhouse temperature between 27 &#176;C and 32 &#176;C to promote optimal adult activity and mating success. The temperature inside the cage is monitored using a traceable memory monitoring thermometer.</li>
	<li>Mist the screened flight cage regularly (approximately once every 2 h) with water using a hand pump, plastic tank sprayer, or garden hose.</li>
	<li>Gently collect individual mating pairs from the screened flight cage using a 50 dram clear plastic snap cap vial (Table of Materials), placing one to two pairs per vial, and transport to an indoor rearing room or laboratory (Supplementary Figure 3).</li>
</ol>
2. Maximizing egg production
<ol>
	<li>Assemble oviposition chamber.
	<ol>
		<li>Take a 12 ounce plain white paper cup and using a snap blade utility knife, make two horizontal cuts on each side of the cup across from one another. Each cut should be approximately 1 cm below the rim.</li>
		<li>Cut a single cotton swab in half and insert the rod end of each into the two horizontal cuts on each side of the paper cup so that the cotton swab portion extends approximately 2 cm towards the interior of the cup.</li>
		<li>Using a snap blade utility knife, make two &#34;X&#34; cuts in the bottom of the paper cup. Each diagonal cut should be approximately 1 cm long.</li>
		<li>Take a 9 ounce plastic cup and fill the bottom with approximately 2 cm of tap water.</li>
		<li>Place a fresh cutting, approximately 15 cm long, of terminal larval host plant growth in the paper cup by inserting the stem through one of the &#34;X&#34; cuts in the bottom. Push the stem through the cut so that approximately 4-5 cm protrudes out the bottom.</li>
		<li>Place the paper cup with host material into the plastic cup, ensuring that the plant stem is in water.</li>
		<li>Fill a 1 ml sub-Q syringe (0.45 mm x 16 mm) with a flavored sports drink and saturate both cotton swabs in the paper cup. These act as artificial flowers. 
		NOTE: Melon and fruit punch flavored sports drink provide the best nectar alternative.</li>
		<li>Once each mating pair separates, place 2-3 gravid females into the assembled cup configuration (i.e., the oviposition chamber).</li>
		<li>Cover the cup with a cut square fragment of black tulle (approximately 15 cm x 15 cm) and secure it with a rubber band around the lid (Supplementary Figure 4). Black tulle provides the best visibility into the cup and easy identification of any eggs that might occasionally be laid on the tulle.</li>
	</ol>
	</li>
	<li>Stimulate adult butterfly activity and oviposition.
	<ol>
		<li>Place each oviposition chamber on a laboratory bench or table approximately 19 cm below an 8.5 inch (21.59 cm) clamp light with an aluminum reflector housing a 40 W incandescent bulb (Supplementary Figure 5). 
		NOTE: The incandescent light provides the radiant heat necessary to stimulate adult activity and oviposition.</li>
		<li>Place a traceable memory monitoring thermometer adjacent to the lights and run the temperature sensor so that it rests on top of an oviposition chamber located directly under a clamp light. 
		NOTE: The target temperature range is between 27.5 &#176;C-29 &#176;C.</li>
		<li>Add supplemental clamp lights as necessary depending on the total number of ovipositional chambers deployed.</li>
		<li>Plug clamp lights into a programmable 15 Amp 24 h indoor plug-in mechanical timer with two outlets (programmable in 30 min timed intervals).</li>
		<li>Set timer to turn clamp light on for 30 min intervals (i.e., a repeatable cycle of 30 min on, 30 min off). 
		NOTE: This light cycle helps to maximize egg production by providing repeatable periods of illumination to stimulate adult butterfly activity and oviposition followed by short dark rest periods.</li>
		<li>Refresh the cotton swabs in each cup with flavored sports drink via the sub-Q syringe and mist regularly with water using a plastic spray bottle approximately every 2-3 h or as needed. 
		NOTE: This provides adequate artificial nectar and moisture to enable the butterflies to free-feed as desired. It thereby enhances both adult longevity and oviposition productivity under laboratory conditions where living, blooming plant material cannot easily be utilized.</li>
		<li>Monitor cups regularly and replace host plant with fresh cuttings as necessary.</li>
		<li>When the eggs start to hatch or the density of eggs becomes high, move the female(s) to a new cup with fresh host and begin larvae protocol with neonates.</li>
	</ol>
	</li>
</ol>
3. Larval care and maintenance
<ol>
	<li>Repeat steps 2.3-2.6 to assemble cups for larvae.</li>
	<li>When eggs begin to hatch, move host plant material with eggs and neonate larvae into a newly assembled cup, placing the stem through the second &#34;X&#34; in the bottom ensuring that the plant stem is in water and leaves touch adjacent fresh host cutting.</li>
	<li>When larvae are young (neonate-2 instar), check larval cups daily for freshness of host plant material and presence of mold or excessive frass. 
	NOTE: Daily removal of host material is not recommended when larvae are young because this can result in organism injury due to handling and/or unnecessary waste of fresh host material.</li>
	<li>If host material is wilted or in otherwise poor condition, place another cutting of fresh host material into cup so that it is touching existing foliage and allow larvae to move to the new host on their own.</li>
	<li>Once larvae reach the 3rd instar, replace paper cup and add fresh host material daily.</li>
	<li>Use a small camel hair watercolor paint brush to gently move the larvae from the old host material or cup surface to fresh host material in the new cup.</li>
	<li>Place the old host material into an empty rectangular plastic food storage container.</li>
	<li>Repeat steps 3.5-3.7 daily and until all cups with larvae have been processed.</li>
	<li>When complete, add a small amount of fresh host material on top of the plant waste in the food storage container and loosely place a lid on top. 
	NOTE: This serves as a safeguard in case any larvae are overlooked during daily processing because they will crawl onto the new host material on top of the plant waste and can be removed the next day.</li>
	<li>Maintain cups under laboratory temperature between 25 &#176;C-28 &#176;C for optimal larval activity and development (Supplementary Figure 6). 
	NOTE: To reach optimal rearing temperatures under indoor conditions, it is often necessary to place cups under clamp lights with aluminum reflectors housing 40 W incandescent bulbs. Temperatures can then be actively monitored using a traceable memory monitoring thermometer and the light height adjusted to reach optimal rearing conditions.</li>
</ol>
4. Constructing the pupation chamber
<ol>
	<li>Cut a single face corrugated paper roll into equal sized 3.8 cm x 3.8 cm squares.</li>
	<li>Place one square into a 2 ounce clear plastic portion cup.</li>
	<li>Place the cup onto a clear plastic cup tray (Supplementary Figure 7).</li>
</ol>
5. Preparing larvae for pupation
<ol>
	<li>Identify mature larvae ready to pupate during daily colony processing. 
	NOTE: Such larvae will turn a uniform dull greenish-brown, lose their chevrons, and often wander off the host.</li>
	<li>Gently remove each mature larva with a small camel hair watercolor paint brush or forceps and place one in each pupation chamber.</li>
	<li>Firmly snap the clear plastic lid on the pupation chamber.</li>
	<li>Repeat steps 5.1-5.3 until all larvae ready to pupate have been transferred into pupation chambers adding new plastic trays as necessary (Supplementary Figure 8).</li>
</ol>
6. Maintaining pupae
<ol>
	<li>For each tray of pupation chambers, record the date of first pupation and any other pertinent information needed (i.e., genetic line, experimental trial, etc.).</li>
	<li>Organize trays by date and place in a secure location within the laboratory (Supplementary Figure 8).</li>
	<li>Monitor trays daily for adult eclosion. 
	NOTE: Laboratory conditions such as temperature will strongly influence development time.</li>
	<li>Prior to adult eclosion (typically within 10 days of first pupation), remove the lids from individual pupation chambers and place the tray in a 34.29 cm x 34.29 cm x 60.96 cm collapsible mesh pop-up rearing cage (Supplementary Figure 9). 
	NOTE: Pupae securely attached within the grooves of the corrugated paper squares facilitate successful adult eclosion (Supplementary Figure 10).</li>
	<li>Repeat the entire protocol from step 1.1 for the subsequent captive generation.</li>
</ol>
7. Assessing development time of immature stages and the number of stadia
<ol>
	<li>Place a single larva under a dissecting microscope. Use a small camel hair watercolor paint brush to carefully move and isolate larvae to avoid injury to the organism.</li>
	<li>Dip a single hair of the paintbrush in nontoxic luminous paint (Table of Materials), and carefully put one small drop of paint on the back (dorsum) of the larva. Use a paint color that stands out from the background color and pattern color of the larva (Supplementary Figure 11). Be sure to avoid placing paint on the head of the larva.</li>
	<li>Once the paint dries (about 30 s or so), place each individual larva in its own 2 ounce clear plastic portion cup containing approximately 1&#8211;3 small leaves of fresh terminal host material and write a unique identifier on the cup and lid (Supplementary Figure 12).</li>
	
	<li>Carefully check each larva daily. Remove leaves and set onto white surface. Inspect cup, clear lid, and examine leaves under a dissecting microscope for the presence of larval exuviae (cast skins) and/or head capsules.</li>
	<li>If a larval exuvia is found, remove it from the cup and place it in a microcentrifuge tube labelled with the corresponding cup number and the date (see steps 8.1-8.6. below).</li>
	<li>Repaint larvae after each molt and record molt dates.</li>
	<li>Measure the total body length (head to last abdominal segment) of each larva daily using digital calipers. Take three measurements and record the average of the three, along with the date and time. For early instar larvae, a magnifying glass or dissecting scope should be used when measuring to ensure accurate measurements.</li>
	<li>Return larva to its corresponding plastic portion cup.</li>
	<li>Add fresh host material as needed and remove all frass and old host debris. If mold is found in the cup, dispose and use a new cup. Write the correct unique identifier number on the new cup.</li>
	<li>Repeat steps 7.5-7.9 until all larvae reach their final instar and begin the prepupal stage. When larvae cease feeding, turn a uniform dull greenish-brown color, lose their chevrons, and often wander off the host, minimize disturbing them.</li>
	<li>Place a small piece of corrugated paper in the cup (see step 4.1).</li>
	<li>Once each larva has fully pupated, measure its total length as in step 7.8 above and record the date of pupation. This will be the final molt of each individual.</li>
	<li>Check on pupae daily and record eclosion date and sex of each resulting adult butterfly.</li>
	<li>Measure the wing chord length of each butterfly using digital calipers. Butterflies can be gently held with forceps for measurement. If the butterfly is too active to easily measure, temporarily place it in a refrigerator for 30 s or less and try again.</li>
</ol>
8. Collecting larval exuviae
<ol>
	<li>When a larval exuvia is observed, fill a microcentrifuge tube with 0.2 &#181;l of glycerin. Label the top of the lid and the side with the larva number, date of molt, and head capsule (H.C.). 
	NOTE: The larvae of some lepidopteran larvae regularly consume their exuviae but the head capsule should remain.</li>
	<li>Place the larval exuvia and the associated head capsule in a clear plastic portion cup lid and put a couple of drops of ethanol in it.</li>
	<li>Examine the larval exuvia under a dissecting microscope by placing it in a clear plastic portion cup lid and putting a few drops of ethanol on it. If the larval head capsule is already separated from the exuvia, place a drop of glycerin on the tip of pointed entomological forceps and gently touch the head capsule to the glycerin. Place the head capsule in the associated microcentrifuge tube.</li>
	<li>If the head capsule is still attached to the larval exuvia, use pointed forceps and an insect pin to separate the head capsule from the larval exuvia.</li>
	<li>Once it is separated, use the glycerin technique to pick up the head capsule. If there is too much ethanol, you can use a small paper towel to remove some, but be careful not to accidentally remove the head capsule.</li>
	<li>Place the head capsule in a labelled glycerin-filled vial and close lid tightly.</li>
</ol>

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D., Russell, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Captive+rearing+of+Puget+blue+butterflies+(Icaricia+icarioides+blackmorei)+and+implications+for+conservation.">Captive rearing of Puget blue butterflies (Icaricia icarioides blackmorei) and implications for conservation.</a> Journal of Insect Conservation. 13 (3), 309-313 (2009).</li><li>Frankham, R., Loebel, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modeling+problems+in+conservation+genetics+using+captive+Drosophila+populations:+Rapid+genetic+adaptation+to+captivity.">Modeling problems in conservation genetics using captive Drosophila populations: Rapid genetic adaptation to captivity.</a> Zoo Biology. 11 (5), 333-342 (1992).</li></ol>

The authors have nothing to disclose.

Here, we illustrate the effectiveness of this proven ex situ conservation breeding protocol for mass production of at-risk butterflies, and how it can be adapted for scientific research to help address key behavioral, life history, or ecological data gaps. Increased understanding of mean total development time (egg to adult), mean duration in each life stage, and optimal temperature for mating, for example, were used to help refine the protocol and enhance overall program success. The vast majority of existing protocols detail only organism husbandry methods and do not discuss data collection, scientific research, or the use of such results to help inform and potentially adapt ex situ methods.
This protocol requires daily organism husbandry. Organism health and productivity are maximized by clean rearing conditions, a lack of organism overcrowding, and the availability of high-quality larval host plant material. For the most part, we utilize disposable rearing supplies and containers (e.g., paper and plastic cups), and typically replace them regularly, often daily, and never reuse the material. This is both cost-effective and minimizes the need for more labor-intensive sanitation of materials. Commonly used tools, however, such as entomological forceps, watercolor paint brushes, and small pop-up flight cages, as well as all rearing surfaces such as tabletops and laboratory bench tops are regularly sanitized using a 5% bleach solution. The exact schedule of sanitation is highly dependent on the frequency of use, organism phenology, and other variables, and should be tailored to the specific needs of each ex situ program. We additionally find that white butcher paper is useful to cover all rearing surfaces. It provides an inexpensive, easily deployable clean substrate, and the white background color facilitates sighting of any stray organisms. For daily husbandry, all laboratory personnel should always wear disposable laboratory exam gloves to minimize contamination and safeguard personnel from any potential skin irritation resulting from plant or organism handling. This is particularly critical if any laboratory personnel have household pets that require topical flea treatments. Even a small amount of active ingredient residue can be hazardous to captive livestock.
Additionally, care should be taken to minimize organism overcrowding. Overcrowding of larvae can quickly lead to reduced organism health and even cannibalism in certain taxa, particularly Lycaenidae. Regularly separating larvae to reduce numbers within rearing containers and/or even isolating individual larvae as described in the life history portion of the protocol may be necessary. The ideal numbers per container may vary considerably based on the particular taxon and various ex situ program constraints such as available budget, laboratory facilities, and total number of husbandry personnel. We similarly recommend leaving adequate space between cups housing larvae to minimize the potential of organism movement between containers. Lastly, for larger captive populations, it is strongly recommended to separate stock between one or more laboratory facilities. This safeguarding strategy can help minimize catastrophic loss of the entire population due to disease or other unforeseen impacts.
Larval host plant quality and availability drives livestock production and strongly influences both larval development rates and overall population health. Nonetheless, few published reports or studies highlight this backstage requirement or discuss best nursery practices. Successful ex situ program planning must account for adequate plant quantities, production, and maintenance. As many larvae also require or prefer certain plant parts (e.g., terminal new growth, flower buds and inflorescences, fruit, etc.), effective staging to ensure appropriate plant phenology is required.
Additional considerations include appropriate demographic and genetic management, and the minimization of any potential negative effects of captivity. We recommend the development of a genetic management plan. This may include strategies to include the infusion of new genetic material on regular basis, maximize diversity and prevent close inbreeding, periodically evaluate key organism fitness variables, and monitor genetics at some level to enable comparison to extant populations and check captive stock health. Periodic comparison of the characteristics of captive individuals to individuals from the founding populations is also warranted34,35.
These protocols represent proven best practices. They should be beneficial to a variety of researchers and conservation practitioners that can directly apply or adapt our methods to their own studies and ex situ at-risk butterfly or insect conservation and recovery programs. The specific outlined captive breeding protocol is likely most applicable to programs focused on other Lycaenidae, related taxa, or smaller sized taxa. Nonetheless, numerous components such as those involving securing successful courtship and copulation, adult maintenance with artificial nectar, maximizing oviposition, and general larval care could arguably be more broadly applied or adapted to a wider array of taxa. As mentioned earlier, while protocol flexibility should be emphasized, access to other established methodologies can help provide valuable insight and a viable departure point for adaptation and innovation. The methods presented for assessing various life history characteristics such as larval development time and the number of larval stadia arguably has broad applicability to other conservation breeding programs and at-risk taxa. We encourage others to help address key ecological data gaps when possible and to publish vetted protocols and program outcomes.

A growing list of studies indicates widespread and serious global declines in butterfly populations1,2,3,4,5. This includes the vast majority of at-risk species. Conservation programs designed to mitigate such declines often employ a mix of strategies including population monitoring, habitat management and restoration, scientific research, captive propagation, and organism translocation6. Within the U.S. and its territories alone, a total of 30 butterfly taxa are listed under the Endangered Species Act (ESA) as either threatened or endangered, with 21 of these having approved draft or final recovery plans. For such taxa, more than half of the identified recovery strategies recommend captive propagation or state that captive propagation should be assessed7. The use of ex situ conservation efforts for butterflies has grown considerably in recent years8,9, and has the potential to be a critical tool to aid recovery efforts10. Numerous institutions, organizations, and agencies are currently involved with ex situ efforts for at least 11 ESA-listed butterfly taxa (i.e., Cyclargus thomasi bethunebakeri, Euphydryas editha quino, Euphydryas editha taylori, Heraclides aristodemus, Hesperia dacotae, Lycaeides melissa samuelis, Oarisma poweshiek, Pyrgus ruralis lagunae, and Speryeria zerene hippolyta) and several other at-risk taxa (e.g., Callophrys irus, Euphydryas phaeton, Speyeria idalia, and Eumaeus atala)11. Despite the number of robust and successful efforts, there remains a lack of regular communication across programs and between conservation practitioners involving the exchange of ideas, data, effective methodologies, and results. Such knowledge sharing is essential as it helps minimize duplication of effort, improves overall best practices, and enhances conservation impact. Few published head-starting, captive rearing, breeding, or husbandry protocols are readily available for at-risk butterfly taxa, and those that are often lack sufficient narrative detail and/or illustrations. These often provide mostly summary details with limited step-by-step instructions and accompanying images, making replication challenging or application to other taxa difficult to assess12,13,14,15. Many of the available protocols are limited in some way: they exist only in the gray literature, or in varying levels of detail, age of publication, or as component parts in symposium proceedings, agency/funder reports, or in-house manuals16,17,18,19,20,21,22,23,24.
For most conservation programs, captive propagation is primarily conducted to support conservation translocation, which encompasses reintroduction, reinforcement (i.e., augmentation), and introduction25,26. Such activities are intended to be implemented strategically as a component of the overall recovery strategy in order to help prevent the extinction of a listed species, subspecies, or populations. It should be noted, however, that this is one of several other potential roles that such ex situ programs may serve. These may also include maintaining an insurance (i.e., refugia) population, temporary organism rescue, supporting recovery-related research and/or training, and promoting conservation-related education and awareness efforts27,28. Regardless of whether ex situ programs have a single defined goal or a mix of several, conservation practitioners should maximize opportunities for data collection in order to fill in key information gaps when possible. This is particularly important because the vast majority of at-risk taxa have generally been poorly studied prior to substantial wild population declines. The resulting enhanced knowledge gained on various behavioral, ecological, or life history aspects of the focal taxon can serve to help advance effective species conservation and management29.
Here, we describe in detail the captive propagation protocol that was developed for the federally endangered Miami blue butterfly (Cyclargus thomasi bethunebakeri) (Supplementary Figure 1) as part of a larger conservation and recovery program. In this case, the captive propagation program serves three specific identified roles: 1) an insurance population should the existing wild population be lost, 2) a research population designed to fill in identified ecological and life history knowledge gaps that may help inform recovery and/or management, and 3) to produce viable organisms for conservation translocation into sites within the taxon&#39;s historical range. The resulting protocol has been well vetted and proven, having been utilized and improved for over a decade. Consequently, we feel that the described techniques and methodologies represent a viable model that can be applied to or readily adapted for other ex situ at-risk butterfly programs, particularly those involving Lycaenidae or related taxa. While we do not suggest that the described protocol is superior to others, we feel that there are opportunities for applying some of the methods more broadly to help enhance productivity, care, or efficiency. This is particularly true as much of our breeding is done under indoor laboratory conditions with limited space, similar to the conservation programs involving Euphydryas editha taylori and Speryeria zerene hippolyta17,23. Numerous other protocols often utilize potted material for oviposition or larval rearing, which can sometimes lead to increased complexities related to predator control, environmental control (i.e., humidity, temperature), livestock monitoring, data collection, plant pest issues, and space to name a few21,22. Lastly, the presented protocol outlines the methods for captive breeding. Many other at-risk butterfly conservation programs involve head-starting or captive rearing with the representative protocols reflecting those differences. While often minor, we feel that this helps broaden the existing pool of available information for other programs to review. This is critical, because most ex situ programs represent pioneering efforts to help facilitate the recovery of rare and often poorly studied taxa. Available protocols can serve as an excellent starting point to help provide valuable insight, reduce duplication of effort, and promote innovation. Owing to &#34;the extensive interspecific diversity of butterfly behaviors, life history traits, and ecological requirements combined with often marked differences in program facilities, budgets, practitioner expertise&#34; and other inherent differences, reliance to a single methodology, even for closely related taxa, is often limiting and unwarranted30. Flexibility to refine or develop new protocols tailored to the needs of specific taxa or programs is essential for success and should therefore be emphasized. We additionally describe laboratory techniques for collecting metrics on organism development under captive conditions including the number of larval instars, duration of individual developmental stages, total development time, and larval and pupal length. These techniques have broad applicability for life history studies of Lepidoptera that can be used to refine ex situ protocols or inform field data.

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at-risk butterfly captive propagation programs to enhance life history knowledge and effective ex situ conservation techniques

Improving knowledge of ex situ best practices for at-risk butterflies is important for generating successful conservation and recovery program outcomes. Research on such captive populations can also yield valuable data to address key information gaps about the behavior, life history, and ecology of the target taxa. We describe a protocol for captive propagation of the federally endangered Cyclargus thomasi bethunebakeri that can be used as a model for other at-risk butterfly ex situ programs, especially those in the family Lycaenidae. We further provide a simple and straightforward protocol for recording various life history metrics that can be useful for informing ex situ methodologies as well as adapted for laboratory studies of other lepidoptera.

Watch this Scientific Journal Video about At-Risk Butterfly Captive Propagation Programs to Enhance Life History Knowledge and Effective Ex Situ Conservation Techniques at JoVE.com

At-Risk Butterfly Captive Propagation Programs to Enhance Life History Knowledge and Effective Ex Situ Conservation Techniques

Here, we present protocols for 1) the laboratory captive propagation of the federally endangered Miami blue butterfly (Cyclargus thomasi bethunebakeri), and 2) assessing basic life history information such as immature development time and number of larval stadia. Both methods can be adapted for use with other ex situ conservation programs.

Improving knowledge of ex situ best practices for at-risk butterflies is important for generating successful conservation and recovery program outcomes. ...

at-risk-butterfly-captive-propagation-programs-to-enhance-life

Research

JoVE Journal

Biology

7.0K Views.  University of Florida. Here, we present protocols for 1) the laboratory captive propagation of the federally endangered Miami blue butterfly (Cyclargus thomasi bethunebakeri), and 2) assessing basic life history information such as immature development time and number of larval stadia. Both methods can be adapted for use with other ex situ conservation programs.

Video: At-Risk Butterfly Captive Propagation Programs to Enhance Life History Knowledge and Effective Ex Situ Conservation Techniques

In situ Protocol for Butterfly Pupal Wings Using Riboprobes

Here we present, in video format, a protocol for in situ hybridizations in pupal wings of the butterfly Bicyclus anynana using riboprobes.  In situ hybridizations, a mainstay of developmental biology, are useful to study the spatial and temporal patterns of gene expression in developing tissues at the level of transcription. If antibodies that target the protein products of gene transcription have not yet been developed, and/or there are multiple gene copies of a particular protein in the genome that cannot be differentiated using available antibodies, in situs can be used instead. While an in situ technique for larval wing discs has been available to the butterfly community for several years, the current protocol has been optimized for the larger and more fragile pupal wings.

In order to examine gene expression in the pupal wing tissue of Bicyclus anynana, we present an optimized protocol for in situ hybridizations using riboprobes. We also provide guidelines for the further optimization of this protocol for use in pupal wings of other Lepidopteran species.

Here we present, in video format, a protocol for in situ hybridizations in pupal wings of the butterfly Bicyclus anynana using riboprobes.  In situ ...

A Noninvasive Hair Sampling Technique to Obtain High Quality DNA from Elusive Small Mammals

Noninvasive genetic sampling approaches are becoming increasingly important to study wildlife populations. A number of studies have reported using noninvasive sampling techniques to investigate population genetics and demography of wild populations1. This approach has proven to be especially useful when dealing with rare or elusive species2. While a number of these methods have been developed to sample hair, feces and other biological material from carnivores and medium-sized mammals, they have largely remained untested in elusive small mammals. In this video, we present a novel, inexpensive and noninvasive hair snare targeted at an elusive small mammal, the American pika (Ochotona princeps). We describe the general set-up of the hair snare, which consists of strips of packing tape arranged in a web-like fashion and placed along travelling routes in the pikas&#x2019; habitat. We illustrate the efficiency of the snare at collecting a large quantity of hair that can then be collected and brought back to the lab. We then demonstrate the use of the DNA IQ system (Promega) to isolate DNA and showcase the utility of this method to amplify commonly used molecular markers including nuclear microsatellites, amplified fragment length polymorphisms (AFLPs), mitochondrial sequences (800bp) as well as a molecular sexing marker. Overall, we demonstrate the utility of this novel noninvasive hair snare as a sampling technique for wildlife population biologists. We anticipate that this approach will be applicable to a variety of small mammals, opening up areas of investigation within natural populations, while minimizing impact to study organisms.

We present a noninvasive sampling approach to efficiently collect hair samples from elusive small mammals, as shown for the American pika. We demonstrate the utility of this method by extracting DNA from sampled hair and amplifying several types of molecular markers commonly used in studies of wildlife ecology and conservation.

Noninvasive genetic sampling approaches are becoming increasingly important to study wildlife populations. A number of studies have reported using ...

Simple and Effective Administration and Visualization of Microparticles in the Circulatory System of Small Fishes Using Kidney Injection

The systemic administration of micro-size particles into a living organism can be applied for vasculature visualization, drug and vaccine delivery, implantation of transgenic cells and tiny optical sensors. However, intravenous microinjections into small animals, which are mostly used in biological and veterinary laboratories, are very difficult and require trained personnel. Herein, we demonstrate a robust and efficient method for the introduction of microparticles into the circulatory system of adult zebrafish (Danio rerio) by injection into the fish kidney. To visualize the introduced microparticles in the vasculature, we propose a simple intravital imaging technique in fish gills. In vivo monitoring of the zebrafish blood pH was accomplished using an injected microencapsulated fluorescent probe, SNARF-1, to demonstrate one of the possible applications of the described technique. This article provides a detailed description of the encapsulation of pH-sensitive dye and demonstrates the principles of the quick injection and visualization of the obtained microcapsules for in vivo recording of the fluorescent signal. The proposed method of injection is characterized by a low mortality rate (0-20%) and high efficiency (70-90% success), and it is easy to institute using commonly available equipment. All described procedures can be performed on other small fish species, such as guppies and medaka.

This article demonstrates the principles of a quick, minimally invasive injection of fluorescent microparticles into the circulatory system of small fishes and the in vivo visualization of the microparticles in fish blood.

The systemic administration of micro-size particles into a living organism can be applied for vasculature visualization, drug and vaccine delivery, ...

Reproductive Techniques for Ovarian Monitoring and Control in Amphibians

Ovarian control and monitoring in amphibians require a multi-faceted approach. There are several applications that can successfully induce reproductive behaviors and the acquisition of gametes and embryos for physiological or molecular research. Amphibians contribute to one-quarter to one-third of vertebrate research, and of interest in this context is their contribution to the scientific community&#39;s knowledge of reproductive processes and embryological development. However, most of this knowledge is derived from a small number of species. In recent times, the decimation of amphibians across the globe has required increasing intervention by conservationists. The captive recovery and assurance colonies that continue to emerge in response to the extinction risk make existing research and clinical applications invaluable to the survival and reproduction of amphibians held under human care. The success of any captive population is founded on its health and reproduction and the ability to develop viable offspring that carry forward the most diverse genetic representation of their species. For researchers and veterinarians, the ability to monitor and control ovarian development and health is, therefore, imperative. The focus of this article is to highlight the different assisted reproductive techniques that can be used to monitor and, where appropriate or necessary, control ovarian function in amphibians. Ideally, any reproductive and health issues should be reduced through proper captive husbandry, but, as with any animal, issues of health and reproductive pathologies are inevitable. Non-invasive techniques include behavioral assessments, visual inspection and palpation and morphometric measurements for the calculation of body condition indices and ultrasound. Invasive techniques include hormonal injections, blood sampling, and surgery. Ovarian control can be exercised in a number of ways depending on the application required and species of interest.

The study of amphibian biology provides valuable information on the reproductive, physiological, embryological and developmental processes that drive organisms of many taxonomic groups. Here, we present a comprehensive guide on different methodologies that can be used to study ovarian control and monitoring in amphibians.

Ovarian control and monitoring in amphibians require a multi-faceted approach. There are several applications that can successfully induce reproductive ...

Applications of Spatio-temporal Mapping and Particle Analysis Techniques to Quantify Intracellular Ca2+ Signaling In Situ

Ca2+ imaging of isolated cells or specific types of cells within intact tissues often reveals complex patterns of Ca2+ signaling. This activity requires careful and in-depth analyses and quantification to capture as much information about the underlying events as possible. Spatial, temporal and intensity parameters intrinsic to Ca2+ signals such as frequency, duration, propagation, velocity and amplitude may provide some biological information required for intracellular signalling. High-resolution Ca2+ imaging typically results in the acquisition of large data files that are time consuming to process in terms of translating the imaging information into quantifiable data, and this process can be susceptible to human error and bias. Analysis of Ca2+ signals from cells in situ typically relies on simple intensity measurements from arbitrarily selected regions of interest (ROI) within a field of view (FOV). This approach ignores much of the important signaling information contained in the FOV. Thus, in order to maximize recovery of information from such high-resolution recordings obtained with Ca2+dyes or optogenetic Ca2+ imaging, appropriate spatial and temporal analysis of the Ca2+ signals is required. The protocols outlined in this paper will describe how a high volume of data can be obtained from Ca2+ imaging recordings to facilitate more complete analysis and quantification of Ca2+ signals recorded from cells using a combination of spatiotemporal map (STM)-based analysis and particle-based analysis. The protocols also describe how different patterns of Ca2+ signaling observed in different cell populations in situ can be analyzed appropriately. For illustration, the method will examine Ca2+ signaling in a specialized population of cells in the small intestine, interstitial cells of Cajal (ICC), using GECIs.

Applications of Spatio-temporal Mapping and Particle Analysis Techniques to Quantify Intracellular Ca2+ Signaling In Situ

Genetically encoded Ca2+ indicators (GECIs) have radically changed how in situ Ca2+ imaging is performed. To maximize data recovery from such recordings, appropriate analysis of Ca2+ signals is required. The protocols in this paper facilitate the quantification of Ca2+ signals recorded in situ using spatiotemporal mapping and particle-based analysis.

Ca2+ imaging of isolated cells or specific types of cells within intact tissues often reveals complex patterns of Ca2+ signaling. This activity requires ...

Analysis of Mitochondrial Respiratory Chain Complexes in Cultured Human Cells using Blue Native Polyacrylamide Gel Electrophoresis and Immunoblotting

Mitochondrial respiration is performed by oxidative phosphorylation (OXPHOS) complexes within mitochondria. Internal and environmental factors can perturb the assembly and stability of OXPHOS complexes. This protocol describes the analysis of mitochondrial respiratory chain complexes by blue native polyacrylamide gel electrophoresis (BN-PAGE) in application to cultured human cells. First, mitochondria are extracted from the cells using digitonin, then using lauryl maltoside, the intact OXPHOS complexes are isolated from the mitochondrial membranes. The OXPHOS complexes are then resolved by gradient gel electrophoresis in the presence of the negatively charged dye, Coomassie blue, which prevents protein aggregation and ensures electrophoretic mobility of protein complexes towards the cathode. Finally, the OXPHOS complexes are detected by standard immunoblotting. Thus, BN-PAGE is a convenient and inexpensive technique that can be used to evaluate the assembly of entire OXPHOS complexes, in contrast to the basic SDS-PAGE allowing the study of only individual OXPHOS complex subunits.

This protocol demonstrates the analysis of mitochondrial respiratory chain complexes by blue native polyacrylamide gel electrophoresis. Here the method applied to cultured human cells is described.

Mitochondrial respiration is performed by oxidative phosphorylation (OXPHOS) complexes within mitochondria. Internal and environmental factors can perturb ...

Glomerular Outgrowth as an Ex Vivo Assay to Analyze Pathways Involved in Parietal Epithelial Cell Activation

Parietal epithelial cell (PEC) activation is one of the key factors involved in the development and progression of glomerulosclerosis. Inhibition of pathways involved in parietal epithelial cell activation could therefore be a tool to attenuate the progression of glomerular diseases. This article describes a method to culture and analyze parietal epithelial cell outgrowth of encapsulated glomeruli isolated from mouse kidney. After dissecting isolated mouse kidneys, the tissue is minced, and glomeruli are isolated by sieving. Encapsulated glomeruli are collected, and single glomeruli are cultured for 6 days to obtain glomerular outgrowth of parietal epithelial cells. During this period, parietal epithelial cell proliferation and migration can be analyzed by determining the cell number or the surface area of outgrowing cells. This assay can therefore be used as a tool to study the effects of an altered gene expression in transgenic- or knockout-mice or the effects of culture conditions on parietal epithelial cell growth characteristics and signaling. Using this method, important pathways involved in the process of parietal epithelial cell activation and consequently in glomerulosclerosis can be studied.

This article describes a method for culturing and analyzing glomerular parietal epithelial cell outgrowths of encapsulated glomeruli isolated from mouse kidney. This method can be used to study pathways involved in parietal epithelial cell proliferation and migration.

Parietal epithelial cell (PEC) activation is one of the key factors involved in the development and progression of glomerulosclerosis. Inhibition of ...

Nucleofection and In Vivo Propagation of Chicken Eimeria Parasites

Transfection is a technical process through which genetic material, such as DNA and double-stranded RNA, are delivered into cells to modify the gene of interest. Currently, transgenic technology is becoming an indispensable tool for the study of Eimeria, the causative agents of coccidiosis in poultry and livestock. This protocol provides a detailed description of stable transfection in eimerian parasites: purification and nucleofection of sporozoites or second-generation merozoites, and in vivo propagation of transfected parasites. Using this protocol, we achieved transfection in several species of Eimeria. Taken together, nucleofection is a useful tool to facilitate genetic manipulation in eimerian parasites.

Nucleofection and In Vivo Propagation of Chicken Eimeria Parasites

Here, we provided a method to achieve stable transfection of chicken Eimeria parasites by nucleofecting sporozoites or second-generation merozoites. Genetically modified eimerian parasites expressing heterologous antigenic genes could be used as vaccine delivery vehicles.

Transfection is a technical process through which genetic material, such as DNA and double-stranded RNA, are delivered into cells to modify the gene of ...

Fluorescence In Situ Hybridization on DNA Halo Preparations to Reveal Whole Chromosomes, Telomeres and Gene Loci

The genome is associated with several structures inside cell nuclei, in order to regulate its activity and anchor it in specific locations. These structures are collectively known as the nucleoskeleton and include the nuclear lamina, the nucleoli, and nuclear bodies. Although many variants of fluorescence in situ hybridization (FISH) exist to study the genome and its organization, these are often limited by resolution and provide insufficient information on the genome&#39;s association with nuclear structures. The DNA halo method uses high salt concentrations and nonionic detergents to generate DNA loops that remain anchored to structures within nuclei through attachment regions within the genome. Here, soluble nuclear proteins, such as histones, lipids, and DNA not tightly bound to the nuclear matrix, are extracted. This leads to the formation of a halo of unattached DNA surrounding a residual nucleus which itself contains DNA closely associated with internal nuclear structures and extraction-resistant proteins. These extended DNA strands enable increased resolution and can facilitate physical mapping. In combination with FISH, this method has the added advantage of studying genomic interactions with all the structures that the genome is anchored by. This technique, termed HALO-FISH, is highly versatile whereby DNA halos can be coupled with nucleic acid probes to reveal gene loci, whole chromosomes, alpha satellite, telomeres and even RNA. This technique provides an insight into nuclear organization and function in normal&#160;cells and in disease progression such as with cancer.

Combining DNA halo preparations with fluorescence in situ hybridization enables high-resolution analysis of genomic interactions with the nucleoskeleton. Attached genome leads to hybridized fluorescent signals located within the residual extracted nuclei, whereas non-attached genome is in the halo of DNA surrounding the residual nuclei.

The genome is associated with several structures inside cell nuclei, in order to regulate its activity and anchor it in specific locations. These ...

Visualizing Scar Development Using SCAD Assay - An Ex-situ Skin Scarring Assay

The mammalian global response to sealing deep tissue wounds is through scar formation and tissue contraction, mediated by specialized fascia fibroblasts. Despite the clinical significance of scar formation and impaired wound healing, our understanding of fascia fibroblast dynamics in wound healing is cursory due to the lack of relevant assays that enable direct visualization of fibroblast choreography and dynamics in complex environments such as in skin wounds. This paper presents a protocol to generate ex- situ skin scars using SCAD or &#34;SCar-like tissue in A Dish&#34; that emulate the complex environment of skin wounds. In this assay, 2 mm full-thickness skin is excised and cultured upside down in media for 5 days, during which scars and skin contractures develop uniformly. This methodology, coupled with fibroblast-lineage specific transgenic mouse models, enables visualization of individual fibroblast lineages across the entire wound repair process. Overall, this protocol aids&#160;researchers in understanding fundamental processes and mechanisms of wound repair,&#160;directly exploring the effects of modulators on wound healing outcomes.

Visualizing Scar Development Using SCAD Assay - An Ex-situ Skin Scarring Assay

This protocol describes the generation of a skin-fascia explant termed "SCar like tissue in A Dish" or SCAD. This model allows unprecedented visualization of single fibroblasts during scar formation.

The mammalian global response to sealing deep tissue wounds is through scar formation and tissue contraction, mediated by specialized fascia fibroblasts. ...