Name: an optimized protocol for rearing fopius arisanus, a parasitoid of tephritid fruit flies
Uploaded: 2011-07-02T14:00:00
Duration: 12 min
Description: Watch this Scientific Journal Video about An Optimized Protocol for Rearing Fopius arisanus, a Parasitoid of Tephritid Fruit Flies at JoVE.com

We thank Keith Shigeteni for assistance in the insectary and Natasha Sostrom for help with computer graphics. This work was funded by USDA-ARS. Opinions, findings, conclusions or recommendations expressed in this publication are those of the authors and do not necessarily reflect the views of the USDA. USDA is an equal opportunity provider and employer.

1. Prepare host fruit fly eggs for parasitization
<ol>
	<li>Prepare a substrate for fruit fly eggs to be parasitized by preparing agar (alternatively, Gelcarin GP812, FMC Biopolymer, Ewing NJ; a more cost effective option) filled dishes, 10 cm in length per side and 1.5 cm depth. Prepare agar at a concentration of 9 g per liter of water.</li>
	<li>Fill dishes to the rim with liquid agar (approximately 70ml), and allow them to cool and solidify (at least 45 minutes).</li>
	<li>Once the agar blocks are solid, apply a single layer of tissue paper (a single ply of Georgia Pacific 'Preference' batch tissue), covering the top of each block. The tissue should stick easily to the agar because of surface moisture.</li>
	<li>Apply a 0.5 ml volume fruit fly eggs suspended in water to the tissue covered surface of the agar blocks. In our insectary, this is equivalent to about 6000 eggs of Bactrocera dorsalis (12000 eggs per ml).</li>
	<li>Use a clean 2.5 cm wide brush dipped in water to spread the fruit fly eggs evenly over the tissue surface.</li>
	<li>Place the fruit fly eggs under the bottom screen of the parasitoid cages to allow oviposition. Tap the underside of the bottom screen to remove dead parasites which may obstruct the eggs. It is best if the bottom screen does not touch the eggs.</li>
	<li>Allow the parasites to oviposit overnight and into the next day, approximately 21 hours. This is sufficient time for a high percentage of the eggs to be parasitized. Longer exposure risks superparasitization.</li>
</ol>
2. Hatching and pupation of parasitized host fruit flies
<ol>
	<li>Prepare rearing container by placing 3 to 3.5 prepared agar blocks of parasitized eggs per 1.25 liters of fruit fly diet19 with eggs facing upward.</li>
	<li>Place the diet container with eggs on a riser over a 1.5 cm deep layer of fine grade vermiculite or washed sand in a larger pupation container. Cover the pupation container with a lid and tape the edges of the with masking tape. Vermiculite may be reused after sifting (see below).</li>
	<li>Ensure that there is sufficient space between the top of the diet container and the pupation container lid for fruit fly larvae to crawl out and 'pop'20 into the vemiculite below.</li>
	<li>Move pupation containers to a room at 27&deg;C and 80 % relative humidity (RH) for one week. Cover containers with a dark cloth for the first four days then remove the cloth for the remaining days.</li>
	<li>Remove larval diet after one week, once larvae have popped and entered the vermiculite for pupation. If there are larvae in the vermiculite, allow them to pupate overnight.</li>
	<li>Sieve the pupae from the vermiculite. Start with a coarse sieve (approximately 3 mm) then use a hand sieve to remove any remaining clumps of vermiculite. </li>
</ol>
3. Sorting and selection
<ol>
	<li>Pupa containing F. arisanus can be partially segregated from unparasitized fruit flies by size. Collect pupa between 1.65 and 2.26 mm in diameter, as these will be enriched for parasitized flies.12 Puparia which are smaller contain mostly undersized parasitoids, larger pupa contain mostly flies. Note also that for the parasitized pupae the percentage of females correlates positively with size12.</li>
	<li>A variety of methods may be employed to sort pupae by size. We will describe a mechanical method using a custom size sorter21.</li>
	<li>Place pupae in the funnel at the top of a vibration feeder which slowly feeds the size sorter.</li>
	<li>Feed the pupae into the size sorter, which consists of a pair of slightly divergent rolling bars at a 30&deg; angle. The pupae drop individually by size into an array of slots that empty into ten separate containers through a funnel.</li>
	<li>Collect pupae from the cups in the size range of 1.65 to 2.26 mm and place them in an emergence cage for 7 more days until most flies have emerged from the unparasitized pupae. </li>
	<li>Next, use a fan to separate empty puparia from those still containing insects.</li>
</ol>
4. F. arisanus emergence and maintenance
<ol>
	<li>To maintain the colony of F. arisanus use a parasitoid holding cage with a removable glass front, approximately 25 cm in length per side. The cage has 1mm2 screening on the top and two of the sides. The back of the cage is covered in a rubber sheet with a 9 cm hole for access. The bottom of the cage has a cut out covered on the inside with 1-2mm2 screen large enough to allow placement of two agar blocks with fruit fly eggs for parasitoid oviposition.</li>
	<li>The holding cages should also have a tinted portion along the bottom quarter of the glass front to minimize light which can create parasitoid crowding along the front of the cage, resulting in elevated mortality.</li>
	<li>Place approximately 11 g of these selected pupae into plastic containers approximately 9 cm in diameter. This should produce about 600-700 parasites per cage. Size selection should result in about 60% females.</li>
	<li>Attach screened lids to the containers. The lids should be covered with 2 mm2 screening, which will allow adult F. arsianus to leave the containers upon emergence but, will contain any remaining fruit flies.</li>
	<li>Maintain the parasites at 24&deg;C and about 45% RH with a 12:12 photoperiod and good ventilation. If high mortality is observed, moving the cages outdoors for a few hours a day or increasing ventilation may reduce it. Parasites should be ready to oviposit in one additional week and should be productive for two more weeks.</li>
	<li>Streak undiluted spun honey along the top of the cages at least three times per week or whenever the available honey is dry. Apply the spun honey in narrow streaks using a fingertip. Note that if the streaks are too heavy the honey will drip into the cage; if too light the honey will dry out quickly and require frequent reapplication.</li>
	<li>Place agar blocks (10 cm x 10 cm x 4 cm) on the tops of the cages to provide moisture to the parasites. These should be changed two times per week.</li>
</ol>
5. Representative Results:
We report here results from quality control procedures conducted on the USDA-ARS F. arisanus rearing operation in Hilo, Hawaii between January 5 and June 22, 2010. Rearing at the Hilo location on the scale described in this video article and protocol was initiated in August 2009, and initial problems and adjustments with the new location had been mostly resolved by January. Therefore, these data are representative of the results other researchers might obtain starting their colony if they follow the protocol as described and have experience rearing insects generally. During this period we were maintaining a small colony of F. arisanus: 4 parasitoid holding cages produced per week, equivalent to about 3600 F. arisanus.
An initial check of enriched parasitization rates was conducted by taking a 2 g sample of pupae in the size range of 1.65 &#x2013; 2.26 mm (enriched) before an additional week of development in a holding cage (i.e. immediately after the larval diet was removed and the pupae sifted and size sorted. We refer to these as 'early pupae'). During the first three months of 2010 the mean proportion parasites in the enriched sample was 0.46 (SD=0.18). During the following quarter this mean was 0.58 (SD=0.08), reflecting stabilization of the colony and rearing procedures at the new location. Note that the enriched pupae which were not successfully parasitized include those which did not produce emergents and those that produced flies.
After an additional week in the holding cage removal of empty puparia another two gram sample was taken ('late pupae'). Fruit flies represented under 1% of this second sample. The low percentage of fruit flies is partially due to to the earlier emergence of flies compared with parasites, so most flies remained in the holding container.
Finally, Figure 1 gives the yield in pupal mass from January to June 2010. The yields are measured from pupae enriched in the size range of interest, 1.65-2.26 mm. The large mass of 'early pupae' at the beginning of the year indicates a high proportion of fruit flies and unemerged pupae at that time, while adjustments were being made to the new facility.
<img src="/files/ftp_upload/2901/2901fig1.jpg" alt="Figure 1" /> 
Figure 1. Yield of enriched (pupal diameter 1.65-2.26mm) early (immediately after size selection) and 'late' (after holding for one week and removing empty casings) pupae at the ARS-USDA F. ariasnus colony in Hilo, Hawaii, January &#x2013; June 2010

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We have no conflicts of interest to report.

In this protocol and accompanying video article we have described and demonstrated an optimized protocol for rearing F. ariasnus, a parasitoid of Tephritid fruit flies, in a laboratory setting. This protocol has been refined over the years to minimize the amount of labor and specialized equipment needed to maintain a colony. We note that a well established, productive and stable colony of host fruit flies is required for any attempt at rearing parasitoids. In our insectary we use Bactrocera dorsalis as a host fruit fly, but other species have been shown to be competent hosts as well14-17.
Several aspects of F. arisanus colony maintenance remain to be explored. These include the rate of laboratory adaptation in this species10, mechanisms of learning22 and the genetic changes that might occur with adaptation and mechanisms which may be involved in host fruit fly plasticity. 
Over the last ten years there have been multiple examples of the spread of Dacine flies throughout the world, particularly those in the genus Bactrocera: B. dorsalis in French Polynesia, B. carambolae in parts of South America, B. invadens in Africa and B. zonata in Africa and Northern Mediterranean.3,23,18 Testing the effectiveness of biological control agents such as F. arisanus against other Bactrocera species should be a high priority, and it is our hope that application of the methods described in this protocol and accompanying video article, will accelerate research on F. arisanus in a wider variety of locations24. Finally, further research using this method as a starting point may also provide important information to the colonization of novel egg parasitoids25,1

an optimized protocol for rearing fopius arisanus, a parasitoid of tephritid fruit flies

Fopius arisanus (Sonan) is an important parasitoid of Tephritid fruit flies for at least two reasons. First, it is the one of only three opiine parasitoids known to infect the host during the egg stage1. Second, it has a wide range of potential fruit fly hosts. Perhaps due to its life history, F. arisanus has been a successfully used for biological control of fruit flies in multiple tropical regions2-4. One impediment to the wide use of F. arisanus for fruit fly control is that it is difficult to establish a stable laboratory colony5-9. Despite this difficulty, in the 1990s USDA researchers developed a reliable method to maintain laboratory populations of F. arisanus10-12. There is significant interest in F. arisanus biology13,14, especially regarding its ability to colonize a wide variety of Tephritid hosts14-17; interest is especially driven by the alarming spread of Bactrocera fruit fly pests to new continents in the last decade18. Further research on F. arisanus and additional deployments of this species as a biological control agent will benefit from optimizations and improvements of rearing methods. In this protocol and associated video article we describe an optimized method for rearing F. arisanus based on a previously described approach12. The method we describe here allows rearing of F. arisanus in a small scale without the use of fruit, using materials available in tropical regions around the world and with relatively low manual labor requirements.

Watch this Scientific Journal Video about An Optimized Protocol for Rearing Fopius arisanus, a Parasitoid of Tephritid Fruit Flies at JoVE.com

An Optimized Protocol for Rearing Fopius arisanus, a Parasitoid of Tephritid Fruit Flies

Fopius arisanus is an egg-larval parasitoid of Tephritid fruit flies that is successfully used in biological control of these important tropical pests. We describe here an optimized protocol for rearing F. arisanus on a small scale using readily available materials.

An Optimized Protocol for Rearing Fopius arisanus, a Parasitoid of Tephritid Fruit Flies

Fopius arisanus (Sonan) is an important parasitoid of Tephritid fruit flies for at least two reasons. First, it is the one of only three opiine ...

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This protocol describes mosquito rearing in the insectary. The insectary rooms are maintained at 28&deg;C and ~80% humidity, with a 12 hr. day/night ...

Isolation and Biophysical Study of Fruit Cuticles

The cuticle, a hydrophobic protective layer on the aerial parts of terrestrial plants, functions as a versatile defensive barrier to various biotic and abiotic stresses and also regulates water flow from the external environment.1 A biopolyester (cutin) and long-chain fatty acids (waxes) form the principal structural framework of the cuticle; the functional integrity of the cuticular layer depends on the outer 'epicuticular' layer as well as the blend consisting of the cutin biopolymer and 'intracuticular' waxes.2 Herein, we describe a comprehensive protocol to extract waxes exhaustively from commercial tomato (Solanum lycopersicum) fruit cuticles or to remove epicuticular and intracuticular waxes sequentially and selectively from the cuticle composite. The method of Jetter and Sch&auml;ffer (2001) was adapted for the stepwise extraction of epicuticular and intracuticular waxes from the fruit cuticle.3,4 To monitor the process of sequential wax removal, solid-state cross-polarization magic-angle-spinning (CPMAS) 13C NMR spectroscopy was used in parallel with atomic force microscopy (AFM), providing molecular-level structural profiles of the bulk materials complemented by information on the microscale topography and roughness of the cuticular surfaces. To evaluate the cross-linking capabilities of dewaxed cuticles from cultivated wild-type and single-gene mutant tomato fruits, MAS 13C NMR was used to compare the relative proportions of oxygenated aliphatic (CHO and CH2O) chemical moieties.
Exhaustive dewaxing by stepwise Soxhlet extraction with a panel of solvents of varying polarity provides an effective means to isolate wax moieties based on the hydrophobic characteristics of their aliphatic and aromatic constituents, while preserving the chemical structure of the cutin biopolyester. The mechanical extraction of epicuticular waxes and selective removal of intracuticular waxes, when monitored by complementary physical methodologies, provides an unprecedented means to investigate the cuticle assembly: this approach reveals the supramolecular organization and structural integration of various types of waxes, the architecture of the cutin-wax matrix, and the chemical composition of each constituent. In addition, solid-state 13C NMR reveals differences in the relative numbers of CHO and CH2O chemical moieties for wild-type and mutant red ripe tomato fruits. The NMR techniques offer exceptional tools to fingerprint the molecular structure of cuticular materials that are insoluble, amorphous, and chemically heterogeneous. As a noninvasive surface-selective imaging technique, AFM furnishes an effective and direct means to probe the structural organization of the cuticular assembly on the nm-&mu;m length scale.

Aerial plant organs are protected by the cuticle, a supramolecular biopolyester-wax assembly. We present protocols to monitor selective removal of epi- and intracuticular waxes from tomato fruit cuticles on molecular and micro scales by solid-state NMR and atomic force microscopy, respectively, and to assess the cross-linking capacity of engineered cuticular biopolyesters.

The cuticle, a hydrophobic protective layer on the aerial parts of terrestrial plants, functions as a versatile defensive barrier to various biotic and ...

Fruit Volatile Analysis Using an Electronic Nose

Numerous and diverse physiological changes occur during fruit ripening, including the development of a specific volatile blend that characterizes fruit aroma. Maturity at harvest is one of the key factors influencing the flavor quality of fruits and vegetables1. The validation of robust methods that rapidly assess fruit maturity and aroma quality would allow improved management of advanced breeding programs, production practices and postharvest handling. 
Over the last three decades, much research has been conducted to develop so-called electronic noses, which are devices able to rapidly detect odors and flavors2-4. Currently there are several commercially available electronic noses able to perform volatile analysis, based on different technologies. The electronic nose used in our work (zNose, EST, Newbury Park, CA, USA), consists of ultra-fast gas chromatography coupled with a surface acoustic wave sensor (UFGC-SAW). This technology has already been tested for its ability to monitor quality of various commodities, including detection of deterioration in apple5; ripeness and rot evaluation in mango6; aroma profiling of thymus species7; C6 volatile compounds in grape berries8; characterization of vegetable oil9 and detection of adulterants in virgin coconut oil10. 
This system can perform the three major steps of aroma analysis: headspace sampling, separation of volatile compounds, and detection. In about one minute, the output, a chromatogram, is produced and, after a purging cycle, the instrument is ready for further analysis. The results obtained with the zNose can be compared to those of other gas-chromatographic systems by calculation of Kovats Indices (KI). Once the instrument has been tuned with an alkane standard solution, the retention times are automatically converted into KIs. However, slight changes in temperature and flow rate are expected to occur over time, causing retention times to drift. Also, depending on the polarity of the column stationary phase, the reproducibility of KI calculations can vary by several index units11. A series of programs and graphical interfaces were therefore developed to compare calculated KIs among samples in a semi-automated fashion. These programs reduce the time required for chromatogram analysis of large data sets and minimize the potential for misinterpretation of the data when chromatograms are not perfectly aligned.
We present a method for rapid volatile compound analysis in fruit. Sample preparation, data acquisition and handling procedures are also discussed.

A rapid method for volatile compound analysis in fruit is described. The volatile compounds present in the headspace of a homogenate of the sample are rapidly separated and detected with ultra-fast gas chromatography (GC) coupled with a surface acoustic wave (SAW) sensor. A procedure for data handling and analysis is also discussed.

Numerous and diverse physiological changes occur during fruit ripening, including the development of a specific volatile blend that characterizes fruit ...

Optimized Protocol for Retinal Wholemount Preparation for Imaging and Immunohistochemistry

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ring-supported hydrophilized PTFE membrane to provide structural support to both living and fixed tissue during immunohistochemical processing, which allows for the use of a variety of protocols that would otherwise cause damage to the tissue. First, this is demonstrated with bolus loading of fluorescent markers into living retinal tissue. This method allows for quick visualization of targeted structures, while the membrane support maintains tissue integrity during the injection and allows for easy transfer of the preparation for further imaging or processing.
Second, a procedure for antibody staining in tissue fixed with carbodiimide is described. Though paraformaldehyde fixation is more common, carbodiimide fixation provides a superior substrate for the visualization of synaptic proteins. A limitation of carbodiimide is that the resulting fixed tissue is relatively fragile; however, this is overcome with the use of the supporting membrane. Retinal tissue is used to demonstrate these techniques, but they may be applied to any fragile tissue.

The protocol described here is for structural assessment of a wholemount retinal preparation. This includes descriptions of tissue dissection, mounting onto a hydrophilized polytetrafluoroethylene (PTFE) membrane insert, bolus loading with fluorescent markers, and a comparison of fixation with carbodiimide and paraformaldehyde for immunohistochemical analysis of cellular and synaptic components.

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ...

Laboratory Rearing of Stable Flies and Other Muscoid Diptera

Stable flies, Stomoxys calcitrans, are serious pests of livestock, humans, companion animals and wildlife worldwide. During the last 20+ years, changes in agronomic practices resulted in serious outbreaks of stable flies in several countries. These outbreaks disrupted livestock production and human recreation resulting in public demands for increasing research and management efforts for this pest. A simple and inexpensive procedure for rearing stable flies for laboratory studies is presented. The procedure uses locally available diet components, equipment and supplies. The procedure can be adapted for rearing other muscoid flies including face fly (Musca autumnalis), horn fly (Haematobia irritans), and house fly (Musca domestica). The procedure produces stable fly puparia averaging 12.5 mg and ~35% egg to adult survival. Approximately 3000 flies are produced in each pan.

A procedure for rearing stable flies (Stomoxys calcitrans) is presented. The procedure uses locally available materials for diet components, equipment and supplies.

Stable flies, Stomoxys calcitrans, are serious pests of livestock, humans, companion animals and wildlife worldwide. During the last 20+ years, changes in ...

Simple Homemade Tools to Handle Fruit Flies&#8212;Drosophila melanogaster

The fruit fly, Drosophila melanogaster, is widely used both in biological research and biology education. Handling adult flies is common but difficult in practice, as adult flies fly. Demonstrated here is how to make some simple and cost-effective tools to address difficult issues in the handling of Drosophila. Holes in foam stoppers are made and pipette tips or funnels are inserted into the holes. Flies then move only in one direction into the pipette tip/funnel assemblage, allowing efficient control of the transfer of adult Drosophila into or out of a vial. Existing protocols have been modified for cool-anesthetizing flies by chilling in crushed ice and transferring them onto a cold, hard icepack surface. The icepack is covered with a piece of medical gauze that keeps immobilized flies from the condensed water when examined under a stereomicroscope. The flies are finally euthanized for counting and sorting or discarded by microwaving. A bottle-shaped cage has also been developed for collecting eggs, as well as a labor-saving device and accompanying protocol for cleaning glass culture vials.

Simple Homemade Tools to Handle Fruit Flies&amp;#8212;&lt;em&gt;Drosophila melanogaster&lt;/em&gt;

Described here is the use of several homemade tools to transfer, chill, and kill adult Drosophila, as well as to clean glass culture vials and collect eggs. These tools are easy to make and are rather efficient in handling Drosophila.

The fruit fly, Drosophila melanogaster, is widely used both in biological research and biology education. Handling adult flies is common but difficult in ...

Topical Application Bioassay to Quantify Insecticide Toxicity for Mosquitoes and Fruit Flies

The continued use of insecticides for public health and agriculture has led to widespread insecticide resistance and hampering of control methods. Insecticide resistance surveillance of mosquito populations is typically done through Centers for Disease Control and Prevention (CDC) bottle bioassays or World Health Organization (WHO) tube tests. However, these methods can result in a high degree of variability in mortality data due to variable insecticide contact with&#160;the insect, the relatively small numbers of organisms tested, extensive variation in mass between populations, and constantly changing environmental conditions, leading to variable outcomes. This paper presents the topical application bioassay, adapted as a high-throughput phenotypic bioassay for both mosquitoes and fruit flies, to test large numbers of insects along&#160;a range of insecticide concentrations.
This assay 1) ensures consistent treatment and insecticide contact with every organism, 2) produces highly specific dose-response curves that account for differences in average mass between strains and sexes (which is particularly important for field-collected organisms), and 3) allows for the calculation of statistically rigorous median lethal doses (LD50), which are necessary for resistance ratio comparisons-an alternative surveillance approach from diagnostic dose mortality, which is also used for larvicide resistance surveillance. This assay will be a complementary tool for accurately phenotyping mosquito populations and, as illustrated using fruit flies, is easily adaptable for use with other insects. We argue that this assay will help fill the gap between genotypic and phenotypic insecticide resistance in multiple insect species.

We describe the methodology and importance of the topical application bioassay to measure insecticide susceptibility in mosquitoes and fruit flies. The presented assay is high-throughput, utilizes insect mass-thus allowing for calculating a mass-relativized lethal dose instead of concentration-and likely has lower variability than other similar methods.

The continued use of insecticides for public health and agriculture has led to widespread insecticide resistance and hampering of control methods. ...

Methods for Rearing the Parasitoid Ganaspis brasiliensis, a Promising Biological Control Agent for the Invasive Drosophila suzukii

Native to East Asia, the spotted-wing drosophila, Drosophila suzukii (Matsumura) (Diptera: Drosophilidae), has established widely in the Americas, Europe, and parts of Africa over the last decade, becoming a devastating pest of various soft-skinned fruits in its invaded regions. Biological control, especially by means of self-perpetuating and specialized parasitoids, is expected to be a viable option for sustainable area-wide management of this highly mobile and polyphagous pest. Ganaspis brasiliensis Ihering (Hymenoptera: Figitidae) is a larval parasitoid that is widely distributed in East Asia, and has been found to be one of the most effective parasitoids of D. suzukii.
Following rigorous pre-introduction evaluations of its efficacy and potential non-target risks, one of the more host-specific genetic groups of this species (G1 G. brasiliensis) has been approved recently for introduction and field release in the United States and Italy. Another genetic group (G3 G. brasiliensis), which was also commonly found to attack D. suzukii in East Asia, may be considered for introduction in the near future. There is currently enormous interest in rearing G. brasiliensis for research or in mass-production for field release against D. suzukii. This protocol and associated video article describe effective rearing methods for this parasitoid, both on a small scale for research and a large scale for mass-production and field release. These methods may benefit further long-term research and use of this Asian-native parasitoid as a promising biological control agent for this global invasive pest.

Methods for Rearing the Parasitoid Ganaspis brasiliensis, a Promising Biological Control Agent for the Invasive Drosophila suzukii

Ganaspis brasiliensis-a larval parasitoid of Drosophila suzukii (a global invasive fruit crop pest)-has been approved or is considered for introduction into Europe and the United States for biological control of this pest. This article provides protocols for both small-scale and large-scale rearing of this parasitoid.

Native to East Asia, the spotted-wing drosophila, Drosophila suzukii (Matsumura) (Diptera: Drosophilidae), has established widely in the Americas, Europe, ...

Enhancing Survival and Development of Marmosets Using a Hand-Rearing Approach

Hand-Rearing Method for Infant Marmosets

The common marmoset (Callithrix jacchus) is a small and highly social New World monkey with high reproduction rates, which has been proven to be a compelling non-human primate model for biomedical and neuroscience research. Some females give birth to triplets; however, the parents cannot raise all of them. To save these infants, we have developed a hand-rearing method for raising newborn marmosets. In this protocol, we describe the formula of the food, the time for feeding, the configuration of the temperature and humidity, as well as the adaptation of the hand-reared infants to the colony environment. This hand-rearing method significantly increases the survival rate of marmoset infants (without hand-rearing: 45%; with hand-rearing: 86%) and provides the opportunity to study the development of marmoset infants with similar genetic backgrounds raised in different postnatal environments. As the method is practical and easy to use, we anticipate that it could also be applied to other labs working with common marmosets.

Author Spotlight: Marmoset Research - Scope and Challenges 

Here, we describe a hand-rearing method for raising infant marmosets in an animal incubator. This method greatly increases the survival rate of marmoset infants, which provides the opportunity to study the development of marmoset infants with similar genetic backgrounds raised in different postnatal environments.