This work was supported by the National Key R&#38;D Program of China (No. 2022YFD1401700) and by the National Science Foundation of China (No. 32090013 and No. 32072385).

1. Insect rearing
<ol>
	<li>Raise the SBPHs used in this experiment in rice seedlings (Oryza sativa cv. Nipponbare). Plant 20 rice seedlings in an incubator (65 mm x 200 mm), and grow at 25 &#176;C under a 16 h light/8 h dark photoperiod.</li>
</ol>
2. Dissection of the SBPHs for hemolymph collection
<ol>
	<li>Put the SBPHs into a centrifuge tube, and place them in an ice bath for 10-30 min. 
	NOTE: Do not place the SBPHs in the ice bath for less than 10 min, or the insects may revive.</li>
	<li>Place a frozen SBPH on a glass slide (see Table of Materials) under a stereomicroscope (see Table of Materials) with its abdomen facing up. Adjust the focus on the six legs of the insect.</li>
	<li>Prepare two high-precision tweezers (see Table of Materials) with ultra-fine tips. Use one tweezer to press down on the insect&#39;s body to keep it in place, and use the other one to carefully pull one of the legs off the insect. 
	NOTE: For insects with a hard shell, an ophthalmic scalpel can be used for cutting off one of the legs.</li>
	<li>Gently press the chest of the insect with a tweezer to make the hemolymph flow out through the wound. 
	&#8203;NOTE: The hemolymph presents as transparent droplets without any visible white floccule. Discard the lipid if there is any white floccule, which represents fat body contamination.</li>
</ol>
3. Hemolymph collection using micropipettes
<ol>
	<li>Prepare a micropipette. Place a capillary tube (see Table of Materials) with a volume of 1 &#181;L into the pipette bulb (see Table of Materials). For accurate measurements, ensure that the scale line is visible. 
	NOTE: A small hole present on the top of the pipette bulb is necessary.</li>
	<li>When collecting the hemolymph, hold the pipette bulb of the micropipette, and place the capillary tube close to the insect wound. Collect the hemolymph exuding from the wound by simply touching the tip of the capillary to the hemolymph.</li>
	<li>Block the small hole on the top of the pipette bulb slightly with the finger to stop the absorbing process when the liquid inside the capillary tube reaches the desired scale line. 
	NOTE: Collect one sample of 1 &#181;L of hemolymph incessantly and rapidly to avoid lipid contamination and tube plugging.</li>
	<li>Press the pipette bulb to discharge the collected hemolymph into 100 &#181;L of PBS buffer (137 mmol/L NaCI, 2.7 mmol/L KCI, 4.3 mmol/L Na2HPO4, 1.4 mmol/L KH2PO4, pH 7.2-7.4). 
	&#8203;NOTE: The capillary tube is inserted into the PBS buffer.</li>
	<li>Change to a new capillary tube when collecting another 1 &#181;L sample of hemolymph.</li>
</ol>
4. Coomassie Blue staining
<ol>
	<li>Collect 3 hemolymph samples with the same volume (1 &#181;L) from 3rd stage larvae according to the protocols described in step 3, and discharge the collected hemolymph into the PBS buffer to make a total volume of 100 &#181;L.</li>
	<li>Add 25 &#181;L of 5x protein loading buffer (250 mmol/L Tris-HCI [pH 6.8], 10% W/V SDS, 0.05% W/V Bromophenol Blue, 50% W/V glycerol, 5% W/V &#946;-mercaptoethanol) into 100 &#181;L of the diluted hemolymph samples, and heat to denature the proteins.</li>
	<li>Load 20 &#181;L of the boiled samples onto a 10% SDS-PAGE protein gel (see Table of Materials) to separate the proteins.</li>
	<li>Stain the gel in a Coomassie Blue solution (mix 1 g of Coomassie Brilliant Blue R-250 with 250 mL of isopropanol, 100 mL of glacial acetic acid, and 650 mL of ddH2O, and filter out any particles using filter paper) for 1 h at room temperature, and then decolorize the gel for 1 h with decolorizing solution (100 mL of acetic acid, 400 mL of methanol, 500 mL of ddH2O).</li>
</ol>
5. Protein concentration determination
<ol>
	<li>Dilute 10 mg/mL bovine serum albumin (BSA) to 5 mg/mL, 2.5 mg/mL, 1.25 mg/mL, 0.625 mg/mL, and 0.3125 mg/mL with PBS buffer using the two-fold dilution method.</li>
	<li>Aspirate 5 &#181;L of PBS buffer and each concentration of BSA solutions in step 5.1, add 1 mL of 1x Bradford dye reagent (see Table of Materials), and incubate for 5 min at room temperature. Use PBS buffer in Bradford dye reagent as the control, and zero at OD = 595 nm. Measure the absorbance values of each of the remaining protein concentrations, and make a standard curve.</li>
	<li>Add 1 &#181;L of hemolymph from larval, female, or male SBPHs into 100 &#181;L of PBS buffer, and then centrifugate the samples at 1,000 x g for 10 min at 4 &#176;C to remove the hemocytes.</li>
	<li>Aspirate 90 &#181;L of supernatant to a new centrifuge tube, and measure the absorbance values of the supernatant according to the instructions in step 5.2.</li>
	<li>Calculate the protein concentrations of the hemolymph samples according to the regression equation of the standard curve. Include three biological replicates with three technical replicates in the experiments.</li>
</ol>
6. Microscopic detections
<ol>
	<li>Use micropipettes to collect hemolymph from 10-15 SBPHs at different developmental stages. Add the collected hemolymph into a tube containing 20 &#181;L of 4% paraformaldehyde solution (see Table of Materials). Incubate the mixture at room temperature for 30 min to fix the hemolymph.</li>
	<li>Pipette 20 &#181;L of fixed hemocytes onto the center of a slide coated with silane (see Table of Materials).</li>
	<li>Dry up the droplet in a slide drying rack. 
	NOTE: Avoid excessive drying.</li>
	<li>Cover the sample with gold antifade reagent 4&#39;,6-diamidino-2-phenylindole (DAPI) (see Table of Materials), and inspect the slide under an inverted microscope (see Table of Materials).</li>
</ol>
7. Cell quantification
<ol>
	<li>Collect 1 &#181;L of hemolymph from the SBPHs with a micropipette, and dissolve it into 20 &#181;L of PBS buffer.</li>
	<li>Dilute the hemolymph samples five-fold with PBS buffer.</li>
	<li>Pipette 20 &#181;L of hemolymph solution into the center of the cell counting chamber (see Table of Materials). Cover the droplet with a coverslip (see Table of Materials).</li>
	<li>Count the cells in the five squares on the cell counting chamber (Supplementary Figure 1), under an inverted microscope (see Table of Materials). Include three biological replicates with three technical replicates in each of the experiments. 
	&#8203;Cells/&#181;L = total count of five middle squares &#215; 5 &#215; dilution factor &#215; 104</li>
</ol>
8. Statistical analyses
<ol>
	<li>Perform statistical analyses with the corresponding software (see Table of Materials). Create a new file, and select the column, import the data, and generate a scatter plot with a bar.</li>
	<li>Calculate the mean and SD for each group of data by using column statistics, and use a one-way ANOVA tool to evaluate the significance of the differences among the groups. Determine the mean and SD from three biological replicates with three independent experiments.</li>
</ol>

Collecting Hemolymph from Small Arthropods

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</ol>

The authors declare that they have no conflicts of interest.

Hemolymph is the medium of the circulatory system in arthropods, and arboviruses can only invade other arthropod tissues if they are able to survive the hostile hemolymph environment. Collecting a high-quality sample of hemolymph is the first step in studying the vector-virus interactions that occur in the hemolymph. It has been reported that insect hemolymph can be obtained from several sites on the insect&#39;s body, including a wound on the front leg, a minor incision in the head area, or a tear wound at the abdomen26,27,28,29. Different collection methods may work for certain acquisitions. The method involving creating a small incision at the head-neck joint is more suitable for hemolymph collection from large insects such as honeybees26. Creating a wound in the head area of the SBPH is challenging as it may damage other organs and introduce contamination from other tissues. After making a tear wound at the abdomen, the insects are immersed in the buffer, and then the buffer is centrifuged to extract the hemolymph. This is a convenient method to collect lipids from small insects21. However, as there is a high amount of fat body within the hemocoel of SBPHs, a wound on the body part may cause fat body leakage from the wound and lead to the contamination of the hemolymph. This method is more suitable for collecting hemolymph from adult SBPHs, which have less fat body than larvae. To effectively prevent fat body contamination, it is recommended to make a wound at the leg site rather than directly on the body. In previous studies of SBPH hemolymph, fine-tipped tweezers were used to facilitate the collection of small drops of liquid from the wound13. Although the collected hemolymph was clear from impurities, the volume of hemolymph was not measured. In this study, we developed a simple micropipette that can be used to accurately and quantitatively collect hemolymph from very small insect vectors.
For successful hemolymph collection, several critical steps must be taken into account. Firstly, it is essential to anesthetize the insects at 4 &#176;C for 15 min in order to produce hemolymph that is free of fat body. Secondly, it is simpler to obtain pure hemolymph by pulling off the first leg. Thirdly, the collection of the hemolymph should be done in a continuous and swift manner, as otherwise the hemolymph may coagulate at the bottom and block the capillary.
The capillary collection technique has been widely used for animal blood collection30,31. We modified the technique to suit the distinct characteristics of insect hemolymph. As SBPHs possess a small body size, a capillary with a minimal volume of 1 &#181;L was chosen. Using a capillary with a volume lower than 1 &#181;L is not recommended. It is noteworthy that hemolymph contains a high density of proteins and a considerable number of cells (Figure 2), which elevate its liquid density and make it difficult to absorb hemolymph via capillaries with smaller volumes.
The method of collecting hemolymph by capillary is a simple, cost-effective, and viable technique that enables the reliable and precise quantification of the hemolymph. This newly developed method could also be applied for collection of other trace fluids from small vectors, such as honeydew. It is vital to highlight that the insects remain alive after the extraction of hemolymph using this method. This is beneficial for studies involving other insect organs or for repeated hemolymph collection. This work presents a valuable technology for the study of entomology and virus-vector interactions.

Both animal and plant viruses can be transmitted by arthropods, and these viruses pose a severe threat to human health and cause tremendous economic losses in agriculture1,2,3. Importantly, the arthropod hemolymph, which serves as the circulatory system and a vital element of the immune system in arthropods, plays an important role in regulating arboviral transmission. Viruses acquired through the arthropod guts are transported to other tissues only after successfully escaping the adverse hemolymph environment4,5,6,7. The lifecycle of viruses in the arthropod hemolymph involves virus survival in the fluid plasma, entry into the hemocyte, and transport to other tissues, and various virus-vector interaction mechanisms occur in the hemolymph8,9,10,11,12. For example, the vertical transmission of RSV by the SBPH is dependent on a molecular interaction between the SBPH vitellogenin protein and the RSV (rice stripe virus) capsid protein13,14. Some viruses may escape the immune response of the hemolymph by binding specific vector factors15,16,17,18. Therefore, investigating vector-virus interactions in the hemolymph of arthropods is important for developing a better understanding of arbovirus transmission.
The hemolymph of some small insects, such as planthoppers, leafhoppers, and some mosquitoes, is difficult to collect due to their size. To address this issue, several methods have been developed to collect hemolymph, including inserting a syringe needle directly into the insect body to extract a microvolume of the hemolymph, collecting exudate from the wound site with fine-tipped tweezers, and direct centrifugation. These methods have enabled the measurement of relative gene expression levels and viral titers within the hemolymph19,20,21. However, an effective method for quantifying the hemolymph volume, which is necessary for hemocyte counting, protein quantification, and enzyme activity analysis, is currently not available for these small insects.
The SBPH (small brown planthopper) is a type of small insect vector with a body length of about 2-4 mm. The SBPH is capable of transmitting a variety of plant viruses, including RSV, maize rough dwarf virus, and rice black streaked dwarf virus22,23,24. The interaction between the SBPH and RSV has been studied in depth over the past decade. To facilitate working with SBPHs, we developed a novel and simple method of collecting hemolymph. This method, which is based on the principle of capillary forces, uses a capillary with a scale mark to acquire the insect&#39;s hemolymph in a precise and quantifiable manner. This allows us to collect a specific volume of hemolymph from small insects efficiently and to study the hemolymph environment of small vectors in more detail.

<table><tbody><tr><td>10% SDS-PAGE protein gel</td><td>Bio-rad</td><td>4561035</td><td>Protein separation and detection</td></tr><tr><td>4% paraformaldehyde</td><td>Solarbio</td><td>P1110</td><td>For fixation of the cells or tissues&amp;nbsp;</td></tr><tr><td>Bradford dye reagent</td><td>Bio-rad</td><td>5000205</td><td>Protein concentration detection</td></tr><tr><td>Capillary</td><td>Hirschmann</td><td>9000101</td><td>For collecting hemolymph</td></tr><tr><td>Cell counting chamber</td><td>ACMEC</td><td>AYA0810</td><td>Hemocytes counting</td></tr><tr><td>Glass slide</td><td>Gitoglas</td><td>10127105A</td><td>For holding insects</td></tr><tr><td>Glass slide coated with silane</td><td>Sigma</td><td>S4651-72EA</td><td>For holding microscope samples</td></tr><tr><td>Gold antifade reagent with DAPI</td><td>Invitrogen</td><td>P36935</td><td>Nucleus staining</td></tr><tr><td>Microscope cover glass</td><td>Gitoglas</td><td>10212424C</td><td>For microscopic observation</td></tr><tr><td>Pipette bulb</td><td>Hirschmann</td><td>9000101</td><td>For collecting hemolymph</td></tr><tr><td>Prism 8.0 software</td><td>GraphPad Software</td><td>/</td><td>Statistical analyses</td></tr><tr><td>Stereomicroscope&amp;nbsp;</td><td>Motic</td><td>SMZ-168</td><td>For insect dissection</td></tr><tr><td>Tweezers</td><td>Tianld</td><td>P5622</td><td>For insect dissection</td></tr><tr><td>Zeiss inverted microscope</td><td>Zeiss</td><td>Observer Z1</td><td>Hemocytes observation</td></tr></tbody></table>

a precise and quantifiable method for collecting hemolymph from small arthropods

Arthropods are known to transmit a variety of viruses of medical and agricultural importance through their hemolymph, which is essential for virus transmission. Hemolymph collection is the basic technology for studying virus-vector interactions. Here, we describe a novel and simple method for the quantitative collection of hemolymph from small arthropods using Laodelphax striatellus (the small brown planthopper, SBPH) as a research model, as this arthropod is the main vector of rice stripe virus (RSV). In this protocol, the process begins by gently pinching off one leg of the frozen arthropod with fine-tipped tweezers and pressing the hemolymph out of the wound. Then, a simple micropipette consisting of a capillary and a pipette bulb is used to collect the transudative hemolymph from the wound according to the principle of capillary forces. Finally, the collected hemolymph can be dissolved into a specific buffer for further study. This new method for collecting hemolymph from small arthropods is a useful and efficient tool for further research on arboviruses and vector-virus interactions.

Micropipette model and hemolymph collection 
We have developed a simple micropipette whose action is based on the capillary forces of the capillary tube. The micropipette is composed of a capillary tube and a pipette bulb (Figure 1A). Capillary tubes are available in different volume sizes ranging from 1 µL to 20 µL, and the capillary tube volumes are selected according to the requirements. Capillary tubes with smaller volumes are not suggested because the extra fine apertures of smaller-volume tubes can make it difficult to absorb liquid such as hemolymph. The pipette bulb contains a hole on the top that cannot be plugged during the hemolymph collection. This pipette bulb is convenient for holding the micropipette during the liquid collection (Figure 1B) and also assists in transferring the collected liquid from the capillary tube into the collection buffer.
In order to collect hemolymph easily, in this work, the SBPHs were firstly frozen in ice or in the refrigerator. These frozen SBPHs were then localized on a slide under a stereomicroscope, and one of the legs of each SBPH was pulled off with fine-tipped tweezers (Figure 1C). To ensure a large wound and optimal hemolymph collection, it is best to pull off the leg at the root (Figure 1Ca, b). In order to minimize the risk of contamination by the fat body, only clear liquid drops without any white floccule were collected (Figure 1Cc). The micropipette was used to absorb the desired volumes of hemolymph (Figure 1Cd). To collect 1 µL of hemolymph, approximately 30-40 larval SBPHs or 8-15 adult SPBHs had to be dissected.
Analysis of the collected hemolymph 
To assess the accuracy of the micropipette as a method for evaluating the volume of collected hemolymph, we tested the protein concentrations of different samples. Hemolymph from larvae was collected using a micropipette with a capillary volume of 1 µL, and three protein samples were collected separately and tested by running an SDS-PAGE gel. The results showed that the amount of protein in the three lanes was nearly equal (Figure 2A). For larvae, the total protein content was 3.707 mg/mL ± 1.382 mg/mL. We also collected the same volume of hemolymph from adult female and male SBPHs, and these showed protein concentrations of 3.515 mg/mL ± 1.400 mg/mL and 3.621 mg/mL ± 0.860 mg/mL, respectively (Table 1). There were no significant differences in the protein among the three samples (Figure 2B).
In addition, we also observed the hemocytes inside the collected larval hemolymph to assess the quality and purity of the hemolymph samples. The hemocytes varied in size between 2-20 nm, and no fat body was detected (Figure 2C). The majority of the cells identified were plasmatocytes, ranging from 5-15 nm in diameter and often appeared in aggregates25. We then counted the cell concentrations of the hemolymph from larvae, adult females, and adult males, and the cell concentrations identified were 1.794 x 105/µL ± 0.614 x 105/µL, 1.256 x 105/µL ± 0.603 x 105/µL, and 1.553 x 105/µL ± 0.474 x 105/µL, respectively (Table 2). The hemocyte cell count in the larvae, adult females, and adult males showed no significant differences (Figure 2D). These results indicate that the micropipette collection method is a reliable and accurate way to collect hemolymph from SBPHs.
<img alt="Capillary injection method diagram; microscopy setup for insect microinjection experiment." class="xfigimg" src="/files/ftp_upload/65250/65250fig01v2.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: Schematic of the micropipette model and hemolymph collection. (A) Micropipette composition. The micropipette consists of a capillary and a pipette bulb. The capillary volumes range from 1-20 µL. The pipette bulb has a small hole on the top and the bottom. The capillary is inserted into the bottom hole, and the scale is visible. (B) Overview diagram of hemolymph collection with a micropipette. The insect abdomen is kept facing up while a leg is detached, hemolymph outflow is induced, and the hemolymph is collected into the pipette under a stereomicroscope. (C) Process of hemolymph collection with a micropipette. One of the legs is pulled off at the root with fine-tipped tweezers (a), and the insect's body is pressed to make hemolymph flow out (b,c). The hemolymph from the wound is collected into the capillary (d). Scale bar = 500 µm. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65250/65250fig01largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Gel electrophoresis protein analysis, hemolymph protein concentration bar chart, cell size microscopy." class="xfigimg" src="/files/ftp_upload/65250/65250fig02.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Analysis of the SBPH hemolymph. (A) Coomassie Blue staining showing three replicates of collected larval hemolymph. Hem indicates hemolymph. (B) Total protein concentrations of the hemolymph of larval, female, and male SBPHs. (C) Microscopic images showing the cells present in the hemolymph in SBPHs at 20x and 60x magnification, respectively. The nucleus was stained with DAPI (blue). Scale bar = 50 µm. (D) Hemocyte density of larval, female, and male SBPHs. The mean and SD were calculated from three independent experiments. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65250/65250fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Hemolymph</td>
			<td>Protein concentration (mg/mL)</td>
		</tr>
		<tr>
			<td>Larva</td>
			<td>3.707±1.382</td>
		</tr>
		<tr>
			<td>Female</td>
			<td>3.515±1.400</td>
		</tr>
		<tr>
			<td>Male</td>
			<td>3.621±0.860</td>
		</tr>
	</tbody>
</table>
Table 1: Protein concentrations of hemolymph from different SBPHs. The data were obtained from three biological replicates.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Hemolymph</td>
			<td>Cell concentration (105 / μL)</td>
		</tr>
		<tr>
			<td>Larva</td>
			<td>1.794±0.614</td>
		</tr>
		<tr>
			<td>Female</td>
			<td>1.256±0.603</td>
		</tr>
		<tr>
			<td>Male</td>
			<td>1.553±0.474</td>
		</tr>
	</tbody>
</table>
Table 2: The total concentrations of hemocytes in the hemolymph from different SBPHs. The data were obtained from three biological replicates.
Supplementary Figure 1: Determining the number of cells. The four corner squares (1, 2, 3, and 4) and the central square (5) are counted on the cell counting chamber. For border cells, only the two boundaries (top and left) are counted. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65250/65250R1_SFigure.pdf" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about A Precise and Quantifiable Method for Collecting Hemolymph from Small Arthropods at JoVE.com

Author Spotlight: A Precise and Quantifiable Method for Collecting Hemolymph from Small Arthropods  

We describe a method to collect quantifiable hemolymph efficiently from small arthropods for subsequent analysis.

A Precise and Quantifiable Method for Collecting Hemolymph from Small Arthropods

Arthropods are known to transmit a variety of viruses of medical and agricultural importance through their hemolymph, which is essential for virus ...

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Universidad San Sebastian

Research

JoVE Journal

Biology

1.6K Views.  Chinese Academy of Sciences. Ons onderzoek richt zich op arbovirussen, insecten en plantinteracties. Het verkrijgen van hemolymfemonsters van kleine geleedpotigen vormt een technische uitdaging voor onderzoek naar door insecten overgedragen ziekten. Dit protocol toont een nauwkeurige en kwantificeerbare techniek om dit probleem tegen te gaan.Recente onderzoeksontwikkelingen in de overdracht van arbovirussen hebben ontdekt dat virussen verschillende belangrijke vectorfactoren kunnen manipuleren om de gastheerbarri?...

Video: Auteur Spotlight: een nauwkeurige en kwantificeerbare methode voor het verzamelen van hemolymfe van kleine geleedpotigen  

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

In vertebrates, hematopoiesis is regulated by inductive microenvironments (niches). Likewise, in the invertebrate model organism&#160;Drosophila melanogaster, inductive microenvironments known as larval Hematopoietic Pockets (HPs) have been identified as anatomical sites for the development and regulation of blood cells (hemocytes), in particular of the self-renewing macrophage lineage.&#160;HPs are segmentally repeated pockets between the epidermis and muscle layers of the larva, which also comprise sensory neurons of the peripheral nervous system. In the larva, resident (sessile) hemocytes are exposed to anti-apoptotic, adhesive and proliferative cues from these sensory neurons and potentially other components of the HPs, such as the lining muscle and epithelial layers. During normal development, gradual release of resident hemocytes from the HPs fuels the population of circulating hemocytes, which culminates in the release of most of the resident hemocytes at the beginning of metamorphosis. Immune assaults, physical injury or mechanical disturbance trigger the premature release of resident hemocytes into circulation. The switch of larval hemocytes between resident locations and circulation raises the need for a common standard/procedure to selectively isolate and quantify these two populations of blood cells from single Drosophila larvae. Accordingly, this protocol describes an automated method to release and quantify the resident and circulating hemocytes from single larvae. The method facilitates ex vivo approaches, and may be adapted to serve a variety of developmental stages of Drosophila and other invertebrate organisms.

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

Drosophila blood cells, or hemocytes, cycle between resident sites and circulation. In the larva, resident (sessile) hemocytes localize to inductive microenvironments, the Hematopoietic Pockets, while circulating hemocytes move freely in the hemolymph. The goal of this protocol is the standardized isolation and quantification of these two, behaviorally distinct but interchanging, hemocyte populations.

In vertebrates, hematopoiesis is regulated by inductive microenvironments (niches). Likewise, in the invertebrate model organism&#160;Drosophila ...

Developmental Biology

A Protocol for Collecting and Staining Hemocytes from the Yellow Fever Mosquito Aedes aegypti

Mosquitoes are vectors for a number of disease-causing pathogens such as the yellow fever virus, malaria parasites and filarial worms. Laboratories are investigating anti-pathogen components of the innate immune system in disease vector species in the hopes of generating transgenic mosquitoes that are refractory to such pathogens1, 2. The innate immune system of mosquitoes consists of several lines of defense 3. Pathogens that manage to escape the barrier imposed by the epithelium-lined mosquito midgut 4 enter the hemolymph and encounter circulating hemocytes, important cellular components that encapsulate and engulf pathogens 5, 6. Researchers have not found evidence for hematopoietic tissues in mosquitoes and current evidence suggests that the number of hemocytes is fixed at adult emergence and numbers may actually decline as the mosquito ages 7. The ability to properly collect and identify hemocytes from medically important insects is an essential step for studies in cellular immunity. However, the small size of mosquitoes and the limited volume of hemolymph pose a challenge to collecting immune cells. 
Two established methods for collecting mosquito hemocytes include expulsion of hemolymph from a cut proboscis 8, and volume displacement (perfusion), in which saline is injected into the membranous necklike region between the head and thorax (i.e., cervix) and the perfused hemolymph is collected from a torn opening in a distal region of the abdomen 9, 10. These techniques, however, are limited by low recovery of hemocytes and possible contamination by fat body cells, respectively 11. More recently a method referred to as high injection/recovery improved recovery of immunocytes by use of anticoagulant buffers while reducing levels of contaminating scales and internal tissues 11. While that method allows for an improved method of collecting and maintaining hemocytes for primary culture, it entails a number of injection and collecting steps that are not necessary if the downstream goal is to collect, fix and stain hemocytes for diagnostics. Here, we demonstrate our method of collecting mosquito hemolymph that combines the simplicity of perfusion, using anticoagulant buffers in place of saline solution, with the accuracy of high injection techniques to isolate clean preparations of hemocytes in Aedes mosquitoes.

A Protocol for Collecting and Staining Hemocytes from the Yellow Fever Mosquito Aedes aegypti

A simplified yet accurate method to collect and stain mosquito hemocytes is described. Our method combines the simplicity of perfusion with the accuracy of high injection techniques to isolate clean preparations of hemocytes in Aedes mosquitoes. This method facilitates studies requiring knowledge of the types of hemocytes and their abundance.

 Mosquitoes are vectors for a number  of disease-causing pathogens such as the yellow fever virus, malaria parasites  and filarial worms. Laboratories are ...

Immunology and Infection

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize mammalian adaptive immunity. Drosophila and mammalian hematopoiesis occur in spatially and temporally distinct phases to produce several blood cell lineages. Both systems maintain reservoirs of blood cell progenitors with which to expand or replace mature lineages. The hematopoietic system allows Drosophila and mammals to respond to and to adapt to immune challenges. Importantly, the transcriptional regulators and signaling pathways that control the generation, maintenance, and function of the hematopoietic system are conserved from flies to mammals. These similarities allow Drosophila to be used to genetically model hematopoietic development and disease.
Here we detail assays to examine the hematopoietic system of Drosophila larvae. In particular, we outline methods to measure blood cell numbers and concentration, visualize a specific mature lineage in vivo, and perform immunohistochemistry on blood cells in circulation and in the hematopoietic organ. These assays can reveal changes in gene expression and cellular processes including signaling, survival, proliferation, and differentiation and can be used to investigate a variety of questions concerning hematopoiesis. Combined with the genetic tools available in Drosophila, these assays can be used to evaluate the hematopoietic system upon defined genetic alterations. While not specifically outlined here, these assays can also be used to examine the effect of environmental alterations, such as infection or diet, on the hematopoietic system.

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Drosophila and mammalian hematopoietic systems share many common features, making Drosophila an attractive genetic model to study hematopoiesis. Here we demonstrate dissection and mounting of the major larval hematopoietic organ for immunohistochemistry. We also describe methods to assay various larval hematopoietic compartments including circulating hemocytes and sessile crystal cells.

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize ...

Harvesting Venom Toxins from Assassin Bugs and Other Heteropteran Insects

Heteropteran insects such as assassin bugs (Reduviidae) and giant water bugs (Belostomatidae) descended from a common predaceous and venomous ancestor, and the majority of extant heteropterans retain this trophic strategy. Some heteropterans have transitioned to feeding on vertebrate blood (such as the kissing bugs, Triatominae; and bed bugs, Cimicidae) while others have reverted to feeding on plants (most Pentatomomorpha). However, with the exception of saliva used by kissing bugs to facilitate blood-feeding, little is known about heteropteran venoms compared to the venoms of spiders, scorpions and snakes.
One obstacle to the characterization of heteropteran venom toxins is the structure and function of the venom/labial glands, which are both morphologically complex and perform multiple biological roles (defense, prey capture, and extra-oral digestion). In this article, we describe three methods we have successfully used to collect heteropteran venoms. First, we present electrostimulation as a convenient way to collect venom that is often lethal when injected into prey animals, and which obviates contamination by glandular tissue. Second, we show that gentle harassment of animals is sufficient to produce venom extrusion from the proboscis and/or venom spitting in some groups of heteropterans. Third, we describe methods to harvest venom toxins by dissection of anaesthetized animals to obtain the venom glands. This method is complementary to other methods, as it may allow harvesting of toxins from taxa in which electrostimulation and harassment are ineffective. These protocols will enable researchers to harvest toxins from heteropteran insects for structure-function characterization and possible applications in medicine and agriculture.

Although many insects in the suborder Heteroptera (Insecta: Hemiptera) are venomous, their venom composition and the functions of their venom toxins are mostly unknown. This protocol describes methods to harvest heteropteran venoms for further characterization, using electrostimulation, harassment, and gland dissection.

Heteropteran insects such as assassin bugs (Reduviidae) and giant water bugs (Belostomatidae) descended from a common predaceous and venomous ancestor, ...

Environment

Insulin Injection and Hemolymph Extraction to Measure Insulin Sensitivity in Adult Drosophila melanogaster

Conserved nutrient sensing mechanisms exist between mammal and fruit fly where peptides resembling mammalian insulin and glucagon, respectively function to maintain glucose homeostasis during developmental larval stages 1,2. Studies on largely post-mitotic adult flies have revealed perturbation of glucose homeostasis as the result of genetic ablation of insulin-like peptide (ILP) producing cells (IPCs) 3. Thus, adult fruit flies hold great promise as a suitable genetic model system for metabolic disorders including type II diabetes. To further develop the fruit fly system, comparable physiological assays used to measure glucose tolerance and insulin sensitivity in mammals must be established. To this end, we have recently described a novel procedure for measuring oral glucose tolerance response in the adult fly and demonstrated the importance of adult IPCs in maintaining glucose homeostasis 4,5. Here, we have modified a previously described procedure for insulin injection 6 and combined it with a novel hemolymph extraction method to measure peripheral insulin sensitivity in the adult fly. Uniquely, our protocol allows direct physiological measurements of the adult fly's ability to dispose of a peripheral glucose load upon insulin injection, a methodology that makes it feasible to characterize insulin signaling mutants and potential interventions affecting glucose tolerance and insulin sensitivity in the adult fly.

Insulin Injection and Hemolymph Extraction to Measure Insulin Sensitivity in Adult Drosophila melanogaster

Conserved insulin signaling pathways found in the fruit fly Drosophila melanogaster make this organism a potential tool for modeling metabolic disorders including type II diabetes. To this end, it is critical to establish physiological assays to effectively measure systemic insulin action in peripheral glucose disposal in the adult fly.

Conserved nutrient sensing mechanisms exist between mammal and fruit fly where peptides resembling mammalian insulin and glucagon, respectively function ...

Extraction of Hemocytes from Drosophila melanogaster Larvae for Microbial Infection and Analysis

During the pathogenic infection of Drosophila melanogaster, hemocytes play an important role in the immune response throughout the infection. Thus, the goal of this protocol is to develop a method to visualize the pathogen invasion in a specific immune compartment of flies, namely hemocytes. Using the method presented here, up to 3 &#215; 106 live hemocytes can be obtained from 200 Drosophila 3rd instar larvae in 30 min for ex vivo infection. Alternatively, hemocytes can be infected in vivo through injection of 3rd instar larvae followed by hemocyte extraction up to 24 h post-infection. These infected primary cells were fixed, stained, and imaged using confocal microscopy. Then, 3D representations were generated from the images to definitively show pathogen invasion. Additionally, high-quality RNA for qRT-PCR can be obtained for the detection of pathogen mRNA following infection, and sufficient protein can be extracted from these cells for Western blot analysis. Taken together, we present a method for definite reconciliation of pathogen invasion and confirmation of infection using bacterial and viral pathogen types and an efficient method for hemocyte extraction to obtain enough live hemocytes from Drosophila larvae for ex vivo and in vivo infection experiments.

Extraction of Hemocytes from Drosophila melanogaster Larvae for Microbial Infection and Analysis

This method demonstrates how to visualize pathogen invasion into insect cells with three-dimensional (3D) models. Hemocytes from Drosophila larvae were infected with viral or bacterial pathogens, either ex vivo or in vivo. Infected hemocytes were then fixed and stained for imaging with a confocal microscope and subsequent 3D cellular reconstruction.

During the pathogenic infection of Drosophila melanogaster, hemocytes play an important role in the immune response throughout the infection. Thus, the ...

A Simple Protocol for Extracting Hemocytes from Wild Caterpillars

Insect hemocytes (equivalent to mammalian white blood cells) play an important role in several physiological processes throughout an insect's life cycle 1. In larval stages of insects belonging to the orders of Lepidoptera (moths and butterflies) and Diptera (true flies), hemocytes are formed from the lymph gland (a specialized hematopoietic organ) or embryonic cells and can be carried through to the adult stage. Embryonic hemocytes are involved in cell migration during development and chemotaxis regulation during inflammation. They also take part in cell apoptosis and are essential for embryogenesis 2. Hemocytes mediate the cellular arm of the insect innate immune response that includes several functions, such as cell spreading, cell aggregation, formation of nodules, phagocytosis and encapsulation of foreign invaders 3. They are also responsible for orchestrating specific insect humoral defenses during infection, such as the production of antimicrobial peptides and other effector molecules 4, 5. Hemocyte morphology and function have mainly been studied in genetic or physiological insect models, including the fruit fly, Drosophila melanogaster 6, 7, the mosquitoes Aedes aegypti and Anopheles gambiae 8, 9 and the tobacco hornworm, Manduca sexta 10, 11. However, little information currently exists about the diversity, classification, morphology and function of hemocytes in non-model insect species, especially those collected from the wild 12.
Here we describe a simple and efficient protocol for extracting hemocytes from wild caterpillars. We use penultimate instar Lithacodes fasciola (yellow-shouldered slug moth) (Figure 1) and Euclea delphinii (spiny oak slug) caterpillars (Lepidoptera: Limacodidae) and show that sufficient volumes of hemolymph (insect blood) can be isolated and hemocyte numbers counted from individual larvae. This method can be used to efficiently study hemocyte types in these species as well as in other related lepidopteran caterpillars harvested from the field, or it can be readily combined with immunological assays designed to investigate hemocyte function following infection with microbial or parasitic organisms 13.

Insect hemocytes carry out many important functions, both immune and non-immune, throughout all stages of insect development. Our present knowledge of hemocyte types and function comes from studies on insect genetic models. Here, we present a method for extracting, quantifying and visualizing hemocytes from wild caterpillars.

Insect hemocytes (equivalent to mammalian white blood cells) play an important role in several physiological processes throughout an insect's life cycle ...

Disclosing Hemolymph Collection and Inoculation of Metarhizium Blastospores into Rhipicephalus Microplus Ticks Towards Invertebrate Pathology Studies

Ticks are obligate hematophagous ectoparasites and Rhipicephalus microplus has great importance in veterinary medicine because it causes anemia, weight loss, depreciation of the animals&#39; leather and also can act as a vector of several pathogens. Due to the exorbitant costs to control these parasites, damage to the environment caused by the inappropriate use of chemical acaricides, and the increased resistance against traditional parasiticides, alternative control of ticks, by the use of entomopathogenic fungi, for example, has been considered an interesting approach. Nevertheless, few studies have demonstrated how the tick&#39;s immune system acts to fight these entomopathogens. Therefore, this protocol demonstrates two methods used for entomopathogen inoculation into engorged females and two techniques used for hemolymph collection and hemocytes harvesting. Inoculation of pathogens at the leg insertion in the tick female&#39;s body allows evaluation of females biologic parameters unlike the inoculation between the scutum and capitulum, which frequently damages Gen&#233;&#39;s organ. Dorsal hemolymph collection yielded a higher volume recovery than collection through the legs. Some limitations of tick hemolymph collection and processing include i) high rates of hemocytes&#39; disruption, ii) hemolymph contamination with disrupted midgut, and iii) low hemolymph volume recovery. When hemolymph is collected through the leg cutting, the hemolymph takes time to accumulate at the leg opening, favoring the clotting process. In addition, fewer hemocytes are obtained in the collection through the leg compared to the dorsal collection, even though the first method is considered easier to be performed. Understanding the immune response in ticks mediated by entomopathogenic agents helps to unveil their pathogenesis and develop new targets for tick control. The inoculation processes described here require very low technological resources and can be used not only to expose ticks to pathogenic microorganisms. Similarly, the collection of tick hemolymph may represent the first step for many physiological studies.

Disclosing Hemolymph Collection and Inoculation of Metarhizium Blastospores into Rhipicephalus Microplus Ticks Towards Invertebrate Pathology Studies

Analysis of tick hemolymph represents an important source of information on how some pathogens cause disease and how ticks immunologically respond to this infection. The present study demonstrates how to inoculate fungal propagules and collect hemolymph from Rhipicephalus microplus engorged females.

Ticks are obligate hematophagous ectoparasites and Rhipicephalus microplus has great importance in veterinary medicine because it causes anemia, weight ...

Extraction of Venom and Venom Gland Microdissections from Spiders for Proteomic and Transcriptomic Analyses

Venoms are chemically complex secretions typically comprising numerous proteins and peptides with varied physiological activities. Functional characterization of venom proteins has important biomedical applications, including the identification of drug leads or probes for cellular receptors. Spiders are the most species rich clade of venomous organisms, but the venoms of only a few species are well-understood, in part due to the difficulty associated with collecting minute quantities of venom from small animals. This paper presents a protocol for the collection of venom from spiders using electrical stimulation, demonstrating the procedure on the Western black widow (Latrodectus hesperus). The collected venom is useful for varied downstream analyses including direct protein identification via mass spectrometry, functional assays, and stimulation of venom gene expression for transcriptomic studies. This technique has the advantage over protocols that isolate venom from whole gland homogenates, which do not separate genuine venom components from cellular proteins that are not secreted as part of the venom. Representative results demonstrate the detection of known venom peptides from the collected sample using mass spectrometry. The venom collection procedure is followed by a protocol for dissecting spider venom glands, with results demonstrating that this leads to the characterization of venom-expressed proteins and peptides at the sequence level.

This article provides a protocol for the extraction of venom from spiders using electrical stimulation in order to 1) conduct proteomic characterization, 2) stimulate venom gland gene expression, and 3) perform functional studies of venoms. This is followed by a description of venom gland microdissections for gene expression studies.

Venoms are chemically complex secretions typically comprising numerous proteins and peptides with varied physiological activities. Functional ...

Cuticle Disruption: A Method to Collect Hemolymph from Drosophila Larvae

Source: Reimels, T. A. and Pfleger, C. M. <a href="https://www-jove-com.bdigitaluss.remotexs.co/video/54544/" target="_blank">Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae</a>. J. Vis. Exp. (2016).
The Drosophila hemolymph serves as both blood and interstitial fluid, circulating throughout the body during the larva and adult stages. This video describes a protocol to collect larval hemolymph in order to quantify the circulating cells.

Source: Reimels, T. A. and Pfleger, C. M. Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae. J. Vis. Exp. (2016).
The Drosophila ...

A Precise and Quantifiable Method for Collecting Hemolymph from Small Arthropods

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