This work was supported by a grant from the Bill &#38; Melinda Gates Foundation and Open Philanthropy. We thank the Facility for Imaging by Light Microscopy (FILM) at Imperial College London for the microscopy analysis. Figure 2 was created with Biorender.com.

1. DNA probe labeling
NOTE: Below are the technical steps to generate fluorescent DNA probes that specifically label the sex chromosomes of An. gambiae mosquitoes.
<ol>
	<li>Probe labeling using PCR
	<ol>
		<li>Extract genomic DNA from pupae or adult males to label the X or Y chromosomes using a commercially available genomic DNA extraction kit (see Table of Materials).</li>
		<li>Prepare the PCR reaction mixture: 200 ng of genomic DNA, 0.05 mM unlabeled nucleotide (dATP, dCTP, dGTP), 0.015 mM of dTTP, and 1 uL of fluorescently labeled dUTP (Cy3, Cy5, or another fluorochrome), 50 pmol of forward and reverse primer (Table 1), 5 &#181;L of 10x PCR-buffer, and 10 U of Taq DNA polymerase (see Table of Materials).</li>
		<li>To label 18S rDNA and satellite AgY53B (Table 1),&#160;perform a PCR reaction using the following PCR parameters: one cycle of 95 &#176;C for 10 min; 35 cycles of 95 &#176;C for 30 s, 52 &#176;C for 30 s, and 72 &#176;C for 45 s; one cycle of 72 &#176;C for 5 min; and a final hold at 4 &#176;C. 
		NOTE: To obtain a good probe concentration with the PCR labeling method (~1 &#181;g in 5 &#181;L), which is critical for a successful WFISH, the PCR reaction should be highly efficient. For this reason, before labeling the probe, testing the efficacy of the primers selected for the amplification is strongly suggested. In addition, including a positive control in the PCR reaction (without the fluorescent dUTP) will help to verify the efficacy of the labeling reaction.</li>
		<li>Store the probe at &#8722;20 &#176;C in a dark place.</li>
	</ol>
	</li>
	<li>Obtaining 3&#39; end fluorescent oligonucleotide probes
	<ol>
		<li>Obtain commercially available fluorescent oligonucleotide probes as modified oligos by adding Cy3 or Cy5 fluorochromes (or any other fluorochrome) to the 3&#39; end of the nucleotide sequence (see Table of Materials). See primers/oligonucleotide in Table 1 for the reference sequences used to label the Y-linked satellite AgY477-AgY53B junction region and the X-linked satellite from Contig_240. 
		&#8203;NOTE: There are no technical impediments related to the concentration of 3&#39; end-labeled oligonucleotides, as the user can usually choose this before purchase. We suggest diluting ~800 ng of oligo probe solution in the hybridization buffer for efficient labeling using the oligo probe. X- and Y-specific oligo-probes have been previously used for WFISH by Liang and Sharakhov19, and the reference sequence can be found in Table 1.</li>
	</ol>
	</li>
</ol>
2. Preparation of the hybridization solution
NOTE: The fluorescent probes generated in step 1 must be incorporated into a chemical solution that hybridizes with the target sequences.
<ol>
	<li>Probe precipitation prior to fluorescence in situ hybridization
	<ol>
		<li>To a 1.5 mL tube, add 5 &#181;L of labeled DNA probe (if obtained by the PCR-labeling method) or 2.2 &#181;L of 3&#39; modified oligo probe (~800 ng of probe) from step 1 and 5 &#181;L of salmon sperm DNA (see Table of Materials). Combine the probes specific to different genomic regions in the same tube, and use them as a unique solution in the following steps.</li>
		<li>Precipitate the DNA probe by adding 0.1 volume of 3 M sodium acetate and 2 volumes of 100% ethanol. Keep at &#8722;20 &#176;C for at least 2.5 h (increasing the time of incubation at &#8722;20 &#176;C will increase the final yield). At this stage, the probes can also be stored overnight before centrifugation.</li>
		<li>Centrifuge at 17,000 x g at 4 &#176;C for 20 min, remove the ethanol, and air-dry the pellet at RT in the dark for ~20 min.</li>
	</ol>
	</li>
	<li>Hybridization solution
	<ol>
		<li>Before proceeding with the testis dissection (step 3), prepare the hybridization buffer by mixing the following reagents in a 1.5 tube: 500 &#181;L of formamide, 0.2 g of dextran sulfate, 100 &#181;L of 20x sodium saline citrate (SSC), and 200 &#181;L of sterile H20 (see Table of Materials). Vortex the hybridization solution for 1 min, and allow the dextran sulfate to dissolve at 37 &#176;C for 30 min.</li>
		<li>Dissolve the pellet from step 2.1.3 in 20-30 &#181;L of hybridization buffer (vortex for about 1 min, perform a quick spin, and store the tubes at 37 &#176;C in the dark) to obtain the hybridization solution.</li>
	</ol>
	</li>
</ol>
3. Testis dissection and fixation
<ol>
	<li>At room temperature (RT), dissect21 at least ~20 testes from pupae or 1 day old adults in sterile 1x phosphate-buffered saline (PBS) solution, and transfer them to a clean microscope slide containing a fresh drop of 1x PBS solution.</li>
	<li>Transfer the testes using a P1,000 wide-bore filtered tip or the tip of a dissection needle from the 1x PBS drop into an embryo dish containing 3.7% formaldehyde in 1x PBS with 0.1% Tween-20 (PBST), and incubate for 10 min at RT.</li>
	<li>Wash the testes in 1x PBST for 5 min at RT. Incubate the testes in 0.1 mg/mL RNAse A (see Table of Materials) diluted in sterile 1x PBS for 30 min at 37 &#176;C.</li>
	<li>Remove the RNAse solution, add the penetrating solution (1% Triton/0.1 M HCl in 1x PBST), and incubate at RT for 10 min. 
	NOTE: Proteinase K can be used as a penetrating agent at a final concentration of 10 &#181;g/mL in 1x PBST to increase the permeabilization of the testes.</li>
	<li>Wash the testes in 1x PBST two times for 5 min each at RT.</li>
</ol>
4. Hybridization
NOTE: This section describes the final steps for the in situ hybridization.
<ol>
	<li>After the washing step (step 3.5), transfer the testes in a 1.5 mL tube with 20-30 &#181;L of hybridization solution containing the previously prepared probe (step 2.2.2). Use the tip of the pipette to mix the solution gently. Gently flick the tube five times before proceeding to the next steps.</li>
	<li>Incubate for 5 min at 75 &#176;C for DNA denaturation.</li>
	<li>Incubate overnight at 37 &#176;C (if possible, with rocking at less than 100 rpm) for DNA-DNA hybridization.</li>
	<li>Transfer the testes back into an embryo dish using a P1,000 wide-bore filtered tip, and wash them in 2x SSC preheated to 50 &#176;C for 5 min. 
	NOTE: After the hybridization step, a final washing step using 2x SSC is required; this plays an important role in removing any background signal caused by the presence of unhybridized probes inside the organ tissue. If a strong background signal is present, repeating the final washing step is recommended.</li>
	<li>Remove the 2x SSC, and mount the testes using a commercially available mounting medium with 4&#8242;,6-diamidino-2-phenylindole (DAPI) (see Table of Materials) on a frosted glass slide, seal with coverslip sealant, and incubate for at least 2 h at RT in the dark.</li>
	<li>Perform confocal imaging. The whole testes can be visualized using 40x or 63x oil immersion objectives. If a z-stack is performed, we suggest using a z-step of 1.25 &#181;m.</li>
</ol>

Fluorescence In Situ Hybridization to Study Mosquito Spermatogenesis

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The authors have nothing to disclose.

Commonly, FISH protocols require the squashing of the organ of interest to allow for chromosome staining. This causes a loss of information regarding the spatial arrangement of the cells within that organ33. This protocol describes how biological processes, such as spermatogenesis, can be studied in situ while maintaining the intact native structure of the testis and its internal cytological organization. Probes targeting different DNA repetitive elements, which are particularly enriched in sex chromosomes20, can be simultaneously used to reveal the dynamics of sperm maturation. Depending on the timing of the testis dissection, WFISH offers the opportunity to study different stages of spermatogenesis through mosquito development. WFISH is useful for studying specific phenomena such as hybrid incompatibility, which, in Anopheles mosquitoes, is due to the presence of meiotic defects such as premeiotic failure and sex chromosome non-disjunction19,34,35. Besides the biological aspect, spermatogenesis is the target of a number of genetic strategies developed to control pest insects such as Anopheles mosquitoes. In this context, the X-linked rDNA locus of An. gambiae has been used as a target to develop a synthetic sex ratio distorter, which, by damaging X-bearing sperms, biases the progeny toward males4,8,13.
This technology mirrors the action of natural sex ratio meiotic drives that have been identified in several taxa, including mosquitoes, but that still remain poorly understood28,36,37,38,39,40,41. WFISH offers the opportunity to investigate this phenomenon and paves the way for refining or improving sex distortion-based genetic strategies by, for example, providing information on how the cytology of sperm production is affected by the choice of the target sites used for sex chromosome shredding. Although, in our experience, WFISH shows high chances of success, failure can still occur. This might be due to an inefficient level of tissue permeabilization, which can be overcome by increasing the incubation time of the penetrating solution. Alternatively, Proteinase K can be used during the permeabilization step. In some cases, we noticed a non-uniform level of probe penetration, with a higher signal in spermatocytes nuclei and a lower or absent signal in the meiotic and spermiogenesis stages. This might be due to a difference in the permeabilization level depending on the cell stage. In addition, WFISH proved to be valuable when using fluorescent probes designed to target DNA sequences present in high copy numbers. When targeting single-copy genes, the signal detection may not be sufficient. In this case, methods for signal amplification, such as tyramide signal amplification (TSA), must be integrated42.
This protocol could be coupled with immunostaining or with transgenic reporter strains harboring germline-specific fluorescent markers16,18, as this would add information about protein localization and gene expression in situ. In this work, WFISH is described as a technique to investigate spermatogenesis in Anopheles mosquitoes; however, given the shared anatomy of male reproductive organs, this protocol could be applied to other mosquito species that play a role in disease transmission. Similarly, female gametogenesis could be investigated using this technique. In addition, cytological studies in organs or tissues of interest, such as the mosquito midgut, which is a target for parasite invasion, or atypical genetic backgrounds, such as those in hybrid mosquitoes, could be explored43. Moreover, this technique can potentially be transferred to other organisms within the Diptera order.

Malaria imposes an enormous burden on the health and well-being of the global human population. In 2021, the World Health Organization (WHO) estimated that malaria caused 619,000 deaths, of which 96% occurred in Sub-Saharan Africa1. The disease is transmitted by mosquitoes belonging to the Anopheles genus, and in Sub-Saharan Africa, three species, namely Anopheles gambiae (An. gambiae), Anopheles coluzzi (An, coluzzi) and Anopheles arabiensis (An. Arabiensis) have a disproportionately large role in malaria transmission, accounting for 95% of malaria cases globally. Control programs relying on traditional methods such as insecticides and antimalarial drugs have saved millions of lives; however, in recent years, the rising resistance to these control methods has challenged their efficacy1,2. In addition, restrictions imposed by the COVID-19 pandemic have affected the availability of key malaria control interventions, which, according to the 2022 WHO World Malaria Report, has increased malaria incidence1. In the last two decades, novel genetic control methods have been developed in laboratory settings to target&#160;Anopheles mosquitoes3,4,5,6,7,8,9,10. Among these strategies, those based on gene drive systems (GDSs) and synthetic sex ratio distorters (SDs) seem promising. GDSs rely on the possibility of transmitting, at a very high frequency, a genetic modification that affects female fertility or impairs the parasite life cycle in the mosquito5,11,12. SDs, instead, act by skewing the sex ratio of a mosquito progeny toward males, which leads, over time, to the collapse of a target population due to a lack of females4,6,13. The main components of these genetic systems act primarily on the reproductive organs of the mosquitoes, where the gametes, eggs, and sperm are produced following meiotic division14.
In this protocol, advances in cytogenetic techniques are employed to explore spermatogenesis in An. gambiae focusing on the chromosomes&#39; behavior in situ. The structure of the mosquito testis and the biological processes that take place within it have been previously investigated using a number of cytological methods, such as immunofluorescence, fluorescent reporter transgenes, and DNA and RNA fluorescence in situ hybridization (FISH)15,16,17,18,19,20; the organs show a spindle-like shape, in which the lower pole is attached to a deferent duct connected to the male accessory glands. In the upper pole, the germline stem cells niche proliferates and differentiates into spermatogonia cells embedded inside spermatocysts formed by somatic cells. Following multiple rounds of mitotic division, the spermatogonia differentiate into spermatocytes, which enter meiosis. At prophase, autosome and sex chromosomes pair with their homologs, and crossing over takes place. After the meiotic divisions, round haploid spermatids are generated and enter spermiogenesis, and this process leads to the formation of mature haploid spermatozoa in which the cytoplasm has been removed, the nuclear chromatin is condensed, and flagella emerge at the basal part of the nuclei21,22 (Figure 1 and Figure 2).
In general, spermiogenesis starts around the mid-pupal stage, and mature spermatozoa can be detected in the late pupal stage in the sperm reservoir23. The maturation process of the spermatocysts continues during adult life23,24,25. In Anopheles testes, each step of spermatogenesis can be easily identified by looking at the nuclear morphology of the cells in each spermatocyst (Figure 2). Whole-mount fluorescence in situ hybridization (WFISH), described in this protocol, allows researchers to specifically label a chromosomal region and track it during spermatogenesis while preserving the native structure of the organ and cell nuclei position; this represents an advantage when compared to the standard DNA FISH protocol in which the organ is usually squashed, leading to tissue damage19. In the current protocol, fluorescent probes are used to stain repetitive sequences on the sex chromosomes and, thus, track their behavior during spermatogenesis, from diploid dividing cells to mature haploid spermatozoa. WFISH can be particularly useful for studying sex chromosome meiotic pairing and investigating the cytological phenotypes associated with, for example, synthetic sex ratio distorters, hybrid male sterility, and the knock-out of genes involved in spermatogenesis4,19,26,27.
Given their role as malaria vectors, Anopheles mosquitoes are the target of an increasing number of genetic vector control strategies, which often act in the reproductive organs of these organisms. Several mosquito mutants and cytological phenotypes have been generated that require novel cytological techniques to be investigated26,27,28,29. The method described in this study sheds light on the understanding of spermatogenesis, as well as the cytological mechanisms behind genetic strategies that have the potential to control malaria-transmitting mosquitoes.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Amersham CyDye Fluorescent Nucleotides, Cy3-dUTP</td>
			<td>Cytiva</td>
			<td>PA53022</td>
			<td></td>
		</tr>
		<tr>
			<td>Amersham CyDye Fluorescent Nucleotides, Cy5-dUTP</td>
			<td>Cytiva</td>
			<td>PA55022</td>
			<td></td>
		</tr>
		<tr>
			<td>ART Wide Bore Filtered Pipette Tips</td>
			<td>ThermoFisher Scientific</td>
			<td>2079GPK</td>
			<td></td>
		</tr>
		<tr>
			<td>CytoBond Removable Coverslip Sealant</td>
			<td>SciGene</td>
			<td>2020-00-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Dextran sulfate sodium salt from Leuconostoc spp.</td>
			<td>Sigma-Aldrich</td>
			<td>D8906-5G</td>
			<td></td>
		</tr>
		<tr>
			<td>DNeasy Blood &#38; Tissue Kits&#160;</td>
			<td>Qiagen</td>
			<td>69504</td>
			<td></td>
		</tr>
		<tr>
			<td>Embryo Dishes</td>
			<td>VWR</td>
			<td>70543-30</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol, molecular grade</td>
			<td>Sigma-Aldrich</td>
			<td>51976</td>
			<td></td>
		</tr>
		<tr>
			<td>Formamide</td>
			<td>ThermoFisher Scientific</td>
			<td>17899</td>
			<td></td>
		</tr>
		<tr>
			<td>GoTaq G2 DNA Polymerase</td>
			<td>Promega</td>
			<td>M7841</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrochloric acid, 37%</td>
			<td>Sigma-Aldrich</td>
			<td>320331</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope slides, SuperFrost</td>
			<td>VWR&#160;</td>
			<td>631-0114</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS (10x), pH 7.4</td>
			<td>ThermoFisher Scientific</td>
			<td>70011044</td>
			<td></td>
		</tr>
		<tr>
			<td>Pierce 16% Formaldehyde (w/v), Methanol-free</td>
			<td>ThermoFisher Scientific</td>
			<td>28906</td>
			<td></td>
		</tr>
		<tr>
			<td>ProLong Gold Antifade Mountant with DAPI</td>
			<td>ThermoFisher Scientific</td>
			<td>P36941</td>
			<td></td>
		</tr>
		<tr>
			<td>RNase A/T1 Mix</td>
			<td>ThermoFisher Scientific</td>
			<td>EN0551</td>
			<td></td>
		</tr>
		<tr>
			<td>Set of dATP, dCTP, dGTP, dTTP</td>
			<td>Promega</td>
			<td>U1330</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Acetate Solution</td>
			<td>ThermoFisher Scientific</td>
			<td>R1181</td>
			<td></td>
		</tr>
		<tr>
			<td>SP8 inverted confocal microscope</td>
			<td>Leica</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma-Aldrich</td>
			<td>9036-19-5</td>
			<td></td>
		</tr>
		<tr>
			<td>TWEEN 20</td>
			<td>Sigma-Aldrich</td>
			<td>P1379</td>
			<td></td>
		</tr>
		<tr>
			<td>UltraPure Salmon Sperm DNA Solution</td>
			<td>ThermoFisher Scientific</td>
			<td>15632011</td>
			<td></td>
		</tr>
		<tr>
			<td>UltraPure SSC 20x</td>
			<td>ThermoFisher Scientific</td>
			<td>15557044</td>
			<td></td>
		</tr>
		<tr>
			<td>Primer sequences</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>5&#8217;-CAATAAATTTCCTTTTTAATGATGC 
			AAAATCTACGTCTCTAGC-3&#8217;-[Fluorochrome]</td>
			<td>Eurofins Genomics</td>
			<td>Contig_240 (X)</td>
			<td></td>
		</tr>
		<tr>
			<td>5&#8217;AGAAGAATAGAATCAGAATAGT 
			CGG 
			TTTCTTCATCCTGAAAGCC-3&#8217;-[Fluorochrome]</td>
			<td>Eurofins Genomics</td>
			<td>AgY53B (Y)</td>
			<td></td>
		</tr>
		<tr>
			<td>5&#8217;-TTCTAAGTTTCTAGGCTTTAAGGA 
			T 
			GAAGAAACCGACTATTC-3&#8217;-[Fluorochrome]</td>
			<td>Eurofins Genomics</td>
			<td>AgY477- 
			AgY53B 
			junction 
			region (Y)</td>
			<td></td>
		</tr>
		<tr>
			<td>F: AACTGTGGAAAAGCCAGAGC 
			R: TCCACTTGATCCTTGCAAAA</td>
			<td>Eurofins Genomics</td>
			<td>18S rDNA (X)</td>
			<td></td>
		</tr>
		<tr>
			<td>F: CCTTTAAACACATGCTCAAATT 
			R: GTTTCTTCATCCTTAAAGCCTAG</td>
			<td>Eurofins Genomics</td>
			<td>AgY53B (Y)</td>
			<td></td>
		</tr>
	</tbody>
</table>

whole-mount fluorescence in situ hybridization to study spermatogenesis in the anopheles mosquito

Spermatogenesis is a complex biological process during which diploid cells undergo successive mitotic and meiotic division followed by large structural changes to form haploid spermatozoa. Besides the biological aspect, studying spermatogenesis is of paramount importance for understanding and developing genetic technologies such as gene drive and synthetic sex ratio distorters, which, by altering Mendelian inheritance and the sperm sex ratio, respectively, could be used to control pest insect populations. These technologies have proven to be very promising in lab settings and could potentially be used to control wild populations of Anopheles mosquitoes, which are vectors of malaria. Due to the simplicity of the testis anatomy and their medical importance, Anopheles gambiae, a major malaria vector in sub-Saharan Africa, represents a good cytological model for studying spermatogenesis. This protocol describes how whole-mount fluorescence in situ hybridization (WFISH) can be used to study the dramatic changes in cell nuclear structure through spermatogenesis using fluorescent probes that specifically stain the X and Y chromosomes. FISH usually requires the disruption of the reproductive organs to expose mitotic or meiotic chromosomes and allow the staining of specific genomic regions with fluorescent probes. WFISH enables the preservation of the native cytological structure of the testis, coupled with a good level of signal detection from fluorescent probes targeting repetitive DNA sequences. This allows researchers to follow changes in the chromosomal behavior of cells undergoing meiosis along the structure of the organ, where each phase of the process can clearly be distinguished. This technique could be particularly useful for studying chromosome meiotic pairing and investigating the cytological phenotypes associated with, for example, synthetic sex ratio distorters, hybrid male sterility, and the knock-out of genes involved in spermatogenesis.

In this work, WFISH was used to investigate the chromosome behavior during spermatogenesis in An. gambiae. The first crucial step for applying this protocol is obtaining testes that show a low level of morphological alteration after dissection. Basic knowledge of the mosquito anatomy is required to perform a successful testis dissection, and below, some guidances for this procedure are given. In the Anopheles mosquito, mature testes can be found lying in the sixth abdominal segment of the pupal and adult stages21. As shown in Figure 1, the vas deferens connects the testes to the male accessory glands (MAGs), which are located in the last segment of the abdomen. The MAGs are connected to a unique ejaculatory duct that delivers the sperms and seminal fluids to the copulatory organ and the external part of the male genital apparatus21. The entire internal male genital apparatus can be dissected using different approaches depending on the life stage of the mosquito. During the pupal stage, testes can be easily identified using a stereomicroscope throughout the light cuticle by looking at the ventral side of the abdomen in the proximity of the sixth segment (Figure 1). To dissect the testes, the lower part of the abdomen, including the sixth segment, can be isolated from the rest of the body using a pair of needles and transferred into a clean drop of 1x PBS. Following the removal of the last segment, the entire apparatus can be squished out of the abdomen by applying gentle pressure with the dissecting needles. To dissect the testes from adult males, the first step involves isolating the entire abdomen in a fresh drop of 1x PBS and then pulling out the last segment carrying the clasps, which are the male copulatory structures (Figure 1). At this point, the lower part of the MAGs should emerge and be easily identifiable due to their yellow color. The entire male apparatus can then be slowly pulled out with the help of a needle or forceps in a drop of 1x PBS until the pair of testes attached to the vas deferens is visible. Before proceeding with the fixation, it is important to isolate the testes from the other parts of the male apparatus by cutting in the proximity of the lower part of the vas deferens (Figure 1).
The age of the pupae or adult males is an important factor to consider depending on the spermatogenesis stage under investigation. In An. gambiae, spermatogenesis starts at the early/mid-pupal stage, and it continues throughout the entire life of the individual24. Between 3 h to 10 h after pupation, the premeiotic and meiotic stages are more represented in the testis (meiotic prophase, meiotic divisions), the spermatid DNA is relatively uncondensed, and mature sperms have not yet been formed. Late pupae and 1 day old adults offer a good balance between the premeiotic, meiotic, and post-meiotic stages (Figure 2 and Figure 3). In adults that are more than 4 days old, the premeiotic stages and spermatocysts are less represented, and the testes are mainly occupied by mature sperms contained in the sperm reservoir.
To investigate the behavior of the sex chromosomes during different stages of spermatogenesis, WFISH was performed on testes dissected at the late pupal stage to ensure a good representation of the entire process. To follow the behavior of these chromosomes, fluorescent probes specific to repetitive sequences located exclusively on the X or Y chromosome were used. The fluorescent probes can be generated using PCR or obtained commercially as 3&#39; end-labeled oligonucleotides. Using an oligo with a length of &#62;40 bps is recommended to allow good signal detection from the fluorescent oligo probe. In our experience, 3&#39; end-labeled oligos perform better than PCR-labeled probes in terms of signal detection. In addition, the copy number of the target sequence is a factor that may affect the efficacy of WFISH. If labeling is unsuccessful, using the PCR labeling method on a longer fragment or designing several oligos specific to the target region is suggested.
The current methods, based on using a penetrating solution (1% Triton/0.1 M HCl in 1x PBST), allow a good level of testis permeabilization and penetration of the probes, resulting in a successful hybridization reaction. Oligo probes specific to sex chromosome repetitive sequences can be designed based on the extensive characterization of repetitive elements performed by Hall et al.20. In addition, consensus sequences specific to X- or Y-linked repetitive elements can be obtained using a bioinformatic platform such as the RedKmer pipeline30. It is important to notice that sex chromosome probes can target repetitive elements such as satellites and retrotransposons, and they can have a different level of hybridization with the X or the Y chromosomes depending on the species under examination20,31,32. As shown in Figure 3, a good level of hybridization of the probes and low background&#160;allowed the visualization of the targeted chromosomes throughout spermatogenesis. The pairing of labeled sex chromosomes could be seen in the premeiotic and meiotic stages. This was followed by detecting either the X or the Y chromosomes in haploid cell nuclei chromosomes resulting from meiotic divisions. Subsequently, X- or Y-bearing spermatids could be followed throughout spermiogenesis, marked by different levels of DNA condensation, to the final step of arrow-shaped mature spermatozoa. In the present experimental setup, confocal Z-stacks were used to acquire information regarding the 3D spatial organization of the cells during this process (Video 1).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65356/65356fig01v2.jpg" /> 
Figure 1: Testes dissected from pupae and 1 day old adult Anopheles gambiae males. (A) An abdomen dissected at the late pupal stage showing the position of the testes in the proximity of the sixth abdominal segment. The testes can be identified throughout the cuticle and appear as brownish structures on both sides of the abdomen (with arrows). (B) Testes dissected at the pupal stage, showing the mature testes (1), vas deferens (2), MAGs (3), and ejaculatory duct (4). (C) An abdomen dissected from a 1 day old adult male after removing the basal clasp segment. The MAGs can be squished from the abdomen by applying gentle pressure (white arrow). (D) Male internal reproductive apparatus dissected from a 1 day old adult male. The white arrow indicates the position occupied by mature sperms, which appear as a white aggregate at the basal pole of the testis. Scale bars: (A,C) 200 &#181;m; (B,D) 100 &#181;m. The Roman numerals from I to VIII indicate the abdominal segments. <a href="https://www.jove.com/files/ftp_upload/65356/65356fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65356/65356fig02v2.jpg" /> 
Figure 2: Representation of spermatogenesis in Anopheles gambiae. The image on the left shows an An. gambiae late pupa testis following whole-mount DAPI staining. On the right is a schematic version for better visualization. By observing the nuclear shape and condensation level, it is relatively easy to follow all the spermatogenesis stages from diploid cells to haploid spermatozoa. The stem cell niche is situated in the upper pole of the organ, where differentiation into spermatogonia starts. The spermatogonia cells increase in number after mitotic division (green cells), and the spermatocysts increase in size (yellow cells). The spermatogonia cells differentiate into spermatocytes after multiple rounds of mitotic divisions (blue cells). The spermatocytes, which are characterized by relatively larger nuclei than the cells of the other stages of the process, are the cells that will undergo meiotic division. Cells undergoing meiosis can be detected by looking at the presence of chromosomes at different meiotic stages; chiasmata and metaphase chromosomes can be detected even at low magnification. The premeiotic stages are overrepresented in testes dissected at the early pupal stage. After the first and second meiotic divisions, spermatids are produced and can usually be found lying in the middle of the testis. The nuclei of spermatids show a certain degree of variation in their shape, from a round to an arrow-like shape. The spermatids enter the spermiogenesis process, during which the nuclei start condensing, and their structure changes into arrow-like dots. When mosquitoes mature sexually after emerging, spermatocysts containing mature sperms can occupy most of the testis volume at the expense of spermatocysts at a different stage of development. Scale bar: 20 &#181;m. The asterisk (*) indicates the apical part of the testis. <a href="https://www.jove.com/files/ftp_upload/65356/65356fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65356/65356fig03v2.jpg" /> 
Figure 3: WFISH on an An. gambiae testis dissected from the late pupal stage. WFISH was performed using probes specific for the X (Contig_240) and Y (oligo probe specific for AgY53B) chromosomes. (A) WFISH allows to&#160;follow&#160;the behavior of the sex chromosome during spermatogenesis from diploid cells to haploid spermatozoa. In this image, it is possible to appreciate the dramatic changes that the nuclei undergo during spermatogenesis. Labeling the sex chromosomes allows for the discrimination between diploid and haploid cells. In diploid cells, the signal from the sex chromosomes is linked to the same nuclei. In haploid cells (spermatids and spermatozoa), the signal of the sex chromosomes is unlinked due to the meiotic reductional division. (B,C) A higher-magnification (63x) image of the testis shown in (A). They were acquired at different positions along the Z-axis. The white dotted frames indicate the area of acquisition. (B) The transition stage between spermatocytes and spermatids, showing the formation of haploid cells and the separation of the signals from the sex chromosomes into separate nuclei. (C) The transition stage between haploid spermatids and mature spermatozoa. This stage shows the changes in the nuclear condensation level; mature spermatozoa show a more condensed and elongated shape than spermatids. Scale bars: (A), 30 &#181;m; (B,C), 10 &#181;m; Gray: DAPI. <a href="https://www.jove.com/files/ftp_upload/65356/65356fig03large.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>Target sequence</td>
			<td>Primer Sequence and Oligo-probe consensus</td>
			<td>Reference</td>
		</tr>
		<tr>
			<td>Contig_240 (X)</td>
			<td>5&#8217;-CAATAAATTTCCTTTTTAATGATGC 
			AAAATCTACGTCTCTAGC-3&#8217;-[Fluorochrome]</td>
			<td>19</td>
		</tr>
		<tr>
			<td>AgY53B (Y)</td>
			<td>5&#8217;AGAAGAATAGAATCAGAATAGTCGG 
			TTTCTTCATCCTGAAAGCC-3&#8217;-[Fluorochrome]</td>
			<td>This study</td>
		</tr>
		<tr>
			<td>AgY477- 
			AgY53B 
			junction 
			region (Y)</td>
			<td>5&#8217;-TTCTAAGTTTCTAGGCTTTAAGGAT 
			GAAGAAACCGACTATTC-3&#8217;-[Fluorochrome]</td>
			<td>19</td>
		</tr>
		<tr>
			<td>18S rDNA (X)</td>
			<td>F: AACTGTGGAAAAGCCAGAGC 
			R: TCCACTTGATCCTTGCAAAA</td>
			<td>19</td>
		</tr>
		<tr>
			<td>AgY53B (Y)</td>
			<td>F: CCTTTAAACACATGCTCAAATT 
			R: GTTTCTTCATCCTTAAAGCCTAG</td>
			<td>19</td>
		</tr>
	</tbody>
</table>
Table 1: List of oligo probes specific to the X or Y chromosome in An. gambiae.
Video 1:&#160;A 3D stack on WFISH performed on an An. gambiae testis dissected from the late pupal stage. To obtain a 3D representation of the spermatogenesis process, a confocal 3D stack can be performed on testes showing a low number of structural alterations. In this study, the stacks were performed with an interval of 1.25 &#181;m between two optical sections under a 63x or 40x oil lens in order not to lose information about the 3D spatial organization of the cells. Gray: DAPI, yellow: Contig_240 (X), magenta: AgY53B (Y). <a href="https://www.jove.com/files/ftp_upload/65356/Video-1.zip" target="_blank">Please click here to download this Video.</a>

Watch this Scientific Journal Video about Whole-Mount Fluorescence In Situ Hybridization to Study Spermatogenesis in the Anopheles Mosquito at JoVE.com

Author Spotlight: Whole-Mount Fluorescence In Situ Hybridization to Study Spermatogenesis in the Anopheles Mosquito  

Given their simple anatomy, Anopheles testes offer a good cytological model for studying spermatogenesis. This protocol describes whole-mount fluorescence in situ hybridization, a technique used to investigate this biological process, as well as the phenotype of transgenic strains harboring mutations in the genes involved in sperm production.

Whole-Mount Fluorescence In Situ Hybridization to Study Spermatogenesis in the Anopheles Mosquito

Spermatogenesis is a complex biological process during which diploid cells undergo successive mitotic and meiotic division followed by large structural ...

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Immunology and Infection

2.2K Views.  Imperial College London. Given their simple anatomy, Anopheles testes offer a good cytological model for studying spermatogenesis. This protocol describes whole-mount fluorescence in situ hybridization, a technique used to investigate this biological process, as well as the phenotype of transgenic strains harboring mutations in the genes involved in sperm production. 

Video: Author Spotlight: Whole-Mount Fluorescence In Situ Hybridization to Study Spermatogenesis in the Anopheles Mosquito  

Combination of Adhesive-tape-based Sampling and Fluorescence in situ Hybridization for Rapid Detection of Salmonella on Fresh Produce

This protocol describes a simple approach for adhesive-tape-based sampling of tomato and other fresh produce surfaces, followed by on-tape fluorescence in situ hybridization (FISH) for rapid culture-independent detection of Salmonella spp. Cell-charged tapes can also be placed face-down on selective agar for solid-phase enrichment prior to detection. Alternatively, low-volume liquid enrichments (liquid surface miniculture) can be performed on the surface of the tape in non-selective broth, followed by FISH and analysis via flow cytometry. To begin, sterile adhesive tape is brought into contact with fresh produce, gentle pressure is applied, and the tape is removed, physically extracting microbes present on these surfaces. Tapes are mounted sticky-side up onto glass microscope slides and the sampled cells are fixed with 10% formalin (30 min) and dehydrated using a graded ethanol series (50, 80, and 95%; 3 min each concentration). Next, cell-charged tapes are spotted with buffer containing a Salmonella-targeted DNA probe cocktail and hybridized for 15 - 30 min at 55&deg;C, followed by a brief rinse in a washing buffer to remove unbound probe. Adherent, FISH-labeled cells are then counterstained with the DNA dye 4',6-diamidino-2-phenylindole (DAPI) and results are viewed using fluorescence microscopy. For solid-phase enrichment, cell-charged tapes are placed face-down on a suitable selective agar surface and incubated to allow in situ growth of Salmonella microcolonies, followed by FISH and microscopy as described above. For liquid surface miniculture, cell-charged tapes are placed sticky side up and a silicone perfusion chamber is applied so that the tape and microscope slide form the bottom of a water-tight chamber into which a small volume (&le; 500 &mu;L) of Trypticase Soy Broth (TSB) is introduced. The inlet ports are sealed and the chambers are incubated at 35 - 37&deg;C, allowing growth-based amplification of tape-extracted microbes. Following incubation, inlet ports are unsealed, cells are detached and mixed with vigorous back and forth pipetting, harvested via centrifugation and fixed in 10% neutral buffered formalin. Finally, samples are hybridized and examined via flow cytometry to reveal the presence of Salmonella spp. As described here, our "tape-FISH" approach can provide simple and rapid sampling and detection of Salmonella on tomato surfaces. We have also used this approach for sampling other types of fresh produce, including spinach and jalape&ntilde;o peppers.

Combination of Adhesive-tape-based Sampling and Fluorescence in situ Hybridization for Rapid Detection of Salmonella on Fresh Produce

This protocol describes a simple adhesive-tape-based approach for sampling of tomato and other fresh produce surfaces, followed by rapid whole cell detection of Salmonella using fluorescence in situ hybridization (FISH).

This protocol describes a simple approach for adhesive-tape-based sampling of tomato and other fresh produce surfaces, followed by on-tape fluorescence in ...

Locked Nucleic Acid Flow Cytometry-fluorescence in situ Hybridization (LNA flow-FISH): a Method for Bacterial Small RNA Detection

Fluorescence in situ hybridization (FISH) is a powerful technique that is used to detect and localize specific nucleic acid sequences in the cellular environment. In order to increase throughput, FISH can be combined with flow cytometry (flow-FISH) to enable the detection of targeted nucleic acid sequences in thousands of individual cells. As a result, flow-FISH offers a distinct advantage over lysate/ensemble-based nucleic acid detection methods because each cell is treated as an independent observation, thereby permitting stronger statistical and variance analyses. These attributes have prompted the use of FISH and flow-FISH methods in a number of different applications and the utility of these methods has been successfully demonstrated in telomere length determination1,2, cellular identification and gene expression3,4, monitoring viral multiplication in infected cells5, and bacterial community analysis and enumeration6.
Traditionally, the specificity of FISH and flow-FISH methods has been imparted by DNA oligonucleotide probes. Recently however, the replacement of DNA oligonucleotide probes with nucleic acid analogs as FISH and flow-FISH probes has increased both the sensitivity and specificity of each technique due to the higher melting temperatures (Tm) of these analogs for natural nucleic acids7,8. Locked nucleic acid (LNA) probes are a type of nucleic acid analog that contain LNA nucleotides spiked throughout a DNA or RNA sequence9,10. When coupled with flow-FISH, LNA probes have previously been shown to outperform conventional DNA probes7,11 and have been successfully used to detect eukaryotic mRNA12 and viral RNA in mammalian cells5.
Here we expand this capability and describe a LNA flow-FISH method which permits the specific detection of RNA in bacterial cells (Figure 1). Specifically, we are interested in the detection of small non-coding regulatory RNA (sRNA) which have garnered considerable interest in the past few years as they have been found to serve as key regulatory elements in many critical cellular processes13. However, there are limited tools to study sRNAs and the challenges of detecting sRNA in bacterial cells is due in part to the relatively small size (typically 50-300 nucleotides in length) and low abundance of sRNA molecules as well as the general difficulty in working with smaller biological cells with varying cellular membranes. In this method, we describe fixation and permeabilzation conditions that preserve the structure of bacterial cells and permit the penetration of LNA probes as well as signal amplification steps which enable the specific detection of low abundance sRNA (Figure 2).

Locked Nucleic Acid Flow Cytometry-fluorescence in situ Hybridization LNA flow-FISH: a Method for Bacterial Small RNA Detection

A novel high-throughput method is described that enables the detection and relative quantitation of small RNA and mRNA expression from single bacterial cells using locked nucleic acid probes and flow cytometry-fluorescence in situ hybridization.

Fluorescence in situ hybridization (FISH) is a powerful technique that is used to detect and localize specific nucleic acid sequences in the cellular ...

Hybridization in situ of Salivary Glands, Ovaries, and Embryos of Vector Mosquitoes

Mosquitoes are vectors for a diverse set of pathogens including arboviruses, protozoan parasites and nematodes. Investigation of transcripts and gene regulators that are expressed in tissues in which the mosquito host and pathogen interact, and in organs involved in reproduction are of great interest for strategies to reduce mosquito-borne disease transmission and disrupt egg development. A number of tools have been employed to study and validate the temporal and tissue-specific regulation of gene expression. Here, we describe protocols that have been developed to obtain spatial information, which enhances our understanding of where specific genes are expressed and their products accumulate. The protocol described has been used to validate expression and determine accumulation patterns of transcripts in tissues related to mosquito-borne pathogen transmission, such as female salivary glands, as well as subcellular compartments of ovaries and embryos, which relate to mosquito reproduction and development. 
 The following procedures represent an optimized methodology that improves the efficiency of various steps in the protocol without loss of target-specific hybridization signals. Guidelines for RNA probe preparation, dissection of soft tissues and the general procedure for fixation and hybridization are described in Part A, while steps specific for the collection, fixation, pre-hybridization and hybridization of mosquito embryos are detailed in Part B.

Hybridization in situ of Salivary Glands, Ovaries, and Embryos of Vector Mosquitoes

Temporal and spatial gene expression analyses have a crucial role in functional genomics. Whole-mount hybridization in situ is useful for determining the localization of transcripts within tissues and subcellular compartments. Here we outline a hybridization in situ protocol with modifications for specific target tissues in mosquitoes.

Mosquitoes are vectors for a diverse set of pathogens including arboviruses, protozoan parasites and nematodes.  Investigation of transcripts and gene ...

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Fluorescent in situ hybridization (FISH) is a technique routinely used by many laboratories to determine the chromosomal position of DNA and RNA probes. One important application of this method is the development of high-quality physical maps useful for improving the genome assemblies for various organisms. The natural banding pattern of polytene and mitotic chromosomes provides guidance for the precise ordering and orientation of the genomic supercontigs. Among the three mosquito genera, namely Anopheles, Aedes, and Culex, a well-established chromosome-based mapping technique has been developed only for Anopheles, whose members possess readable polytene chromosomes 1. As a result of genome mapping efforts, 88% of the An. gambiae genome has been placed to precise chromosome positions 2,3 . Two other mosquito genera, Aedes and Culex, have poorly polytenized chromosomes because of significant overrepresentation of transposable elements in their genomes 4, 5, 6. Only 31 and 9% of the genomic supercontings have been assigned without order or orientation to chromosomes of Ae. aegypti 7 and Cx. quinquefasciatus 8, respectively. Mitotic chromosome preparation for these two species had previously been limited to brain ganglia and cell lines. However, chromosome slides prepared from the brain ganglia of mosquitoes usually contain low numbers of metaphase plates 9. Also, although a FISH technique has been developed for mitotic chromosomes from a cell line of Ae. aegypti 10, the accumulation of multiple chromosomal rearrangements in cell line chromosomes 11 makes them useless for genome mapping. Here we describe a simple, robust technique for obtaining high-quality mitotic chromosome preparations from imaginal discs (IDs) of 4th instar larvae which can be used for all three genera of mosquitoes. A standard FISH protocol 12 is optimized for using BAC clones of genomic DNA as a probe on mitotic chromosomes of Ae. aegypti and Cx. quinquefasciatus, and for utilizing an intergenic spacer (IGS) region of ribosomal DNA (rDNA) as a probe on An. gambiae chromosomes. In addition to physical mapping, the developed technique can be applied to population cytogenetics and chromosome taxonomy/systematics of mosquitoes and other insect groups.

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Among the three mosquito genera, namely Anopheles, Aedes, and Culex, physical genome mapping techniques were established only for Anopheles, whose members possess readable polytene chromosomes. For the genera of Aedes and Culex, however, cytogenetic mapping remains challenging because of the poor quality of polytene chromosomes. Here we present a universal protocol for obtaining high-quality preparations of mitotic chromosomes and an optimized FISH protocol for all three genera of mosquitoes.

Fluorescent in situ hybridization (FISH) is a technique routinely used by many laboratories to determine the chromosomal position of DNA and RNA probes. ...

Fluorescence in situ Hybridizations (FISH) for the Localization of Viruses and Endosymbiotic Bacteria in Plant and Insect Tissues

Fluorescence in situ hybridization (FISH) is a name given to a variety of techniques commonly used for visualizing gene transcripts in eukaryotic cells and can be further modified to visualize other components in the cell such as infection with viruses and bacteria. Spatial localization and visualization of viruses and bacteria during the infection process is an essential step that complements expression profiling experiments such as microarrays and RNAseq in response to different stimuli. Understanding the spatiotemporal infections with these agents complements biological experiments aimed at understanding their interaction with cellular components. Several techniques for visualizing viruses and bacteria such as reporter gene systems or immunohistochemical methods are time-consuming, and some are limited to work with model organisms&#160;and involve complex methodologies. FISH that targets RNA or DNA species in the cell is a relatively easy and fast method for studying spatiotemporal localization of genes and for diagnostic purposes. This method can be robust and relatively easy to implement when the protocols employ short hybridizing, commercially-purchased probes, which are not expensive. This is particularly robust when sample preparation, fixation, hybridization, and microscopic visualization do not involve complex steps. Here we describe a protocol for localization of bacteria and viruses in insect and plant tissues. The method is based on simple preparation, fixation, and hybridization of insect whole mounts and dissected organs or hand-made plant sections, with 20 base pairs short DNA probes conjugated to fluorescent dyes on their 5&#39; or 3&#39; ends. This protocol has been successfully applied to a number of insect and plant tissues, and can be used to analyze expression of mRNAs or other RNA or DNA species in the cell.

Fluorescence in situ Hybridizations FISH for the Localization of Viruses and Endosymbiotic Bacteria in Plant and Insect Tissues

We describe here a simple fluorescence in situ hybridization (FISH) method for the localization of viruses and bacteria in insect and plant tissues. This protocol can be extended for the visualization of mRNA in whole mount and microscopic sections.

Fluorescence in situ hybridization (FISH) is a name given to a variety of techniques commonly used for visualizing gene transcripts in eukaryotic cells ...

Human Placental and Decidual Organ Cultures to Study Infections at the Maternal-fetal Interface

The placenta shows a large degree of interspecies anatomic variability. To best understand biology and pathophysiology of the human placenta, it is imperative to design experiments using human cells and tissues. An advantage of organ culture is maintenance of three-dimensional (3D) structural organization and extracellular matrix. The goal of the method described here is successful establishment of ex vivo human gestational tissue organ cultures and their healthy culture maintenance for 72-96 hr. The protocol details the immediate processing of research-consented, placental and decidual specimens fresh from the operating suite. These are abundant specimens that would otherwise be discarded. Detailed instructions on the sterile collection of these samples, including morphologic details on how to select appropriate tissues to establish 3D organ cultures, is provided. Placental villous and decidual tissues are microdissected into 2-3 mm3 pieces and placed separately on matrix-lined transwell filters and cultured for several days. Villous and decidual organ cultures are well suited for the study of human host-pathogen interaction. As compared to other model organisms, these human cultures are particularly advantageous to examine mechanism of infection for pathogens that demonstrate variable patterns of host specificity. As an example, we demonstrate infection of placental and decidual organ cultures with the clinically relevant, facultative intracellular bacterial pathogen Listeria monocytogenes.

A simple method to establish primary human placental (villous) and decidual organ cultures is described. Villous and decidual organ cultures are invaluable tools for studying pathogenesis at the human maternal-fetal interface. Infection with the facultative intracellular bacterium Listeria monocytogenes is demonstrated.

The placenta shows a large degree of interspecies anatomic variability. To best understand biology and pathophysiology of the human placenta, it is ...

Overexpression and Purification of Human Cis-prenyltransferase in Escherichia coli

Prenyltransferases (PT) are a group of enzymes that catalyze chain elongation of allylic diphosphate using isopentenyl diphosphate (IPP) via multiple condensation reactions. DHDDS (dehydrodolichyl diphosphate synthase) is a eukaryotic long-chain cis-PT (forming cis double bonds from the condensation reaction) that catalyzes chain elongation of farnesyl diphosphate (FPP, an allylic diphosphate) via multiple condensations with isopentenyl diphosphate (IPP). DHDDS is of biomedical importance, as a non-conservative mutation (K42E) in the enzyme results in retinitis pigmentosa, ultimately leading to blindness. Therefore, the present protocol was developed in order to acquire large quantities of purified DHDDS, suitable for mechanistic studies. Here, the usage of protein fusion, optimized culture conditions and codon-optimization were used to allow the overexpression and purification of functionally active human DHDDS in E. coli. The described protocol is simple, cost-effective and time sparing. The homology of cis-PT among different species suggests that this protocol may be applied for other eukaryotic cis-PT as well, such as those involved in natural rubber synthesis.

Overexpression and Purification of Human Cis-prenyltransferase in Escherichia coli

A simple protocol for overexpression and purification of codon-optimized, human cis-prenyltransferase, under non-denaturing conditions, from Escherichia coli, is described, along with an enzymatic activity assay. This protocol can be generalized for production of other cis- prenyltransferase proteins in quantity and quality suitable for mechanistic studies.

Prenyltransferases (PT) are a group of enzymes that catalyze chain elongation of allylic diphosphate using isopentenyl diphosphate (IPP) via multiple ...

Human Colonoid Monolayers to Study Interactions Between Pathogens, Commensals, and Host Intestinal Epithelium

Human 3-dimensional (3D) enteroid or colonoid cultures derived from crypt base stem cells are currently the most advanced ex vivo model of the intestinal epithelium. Due to their closed structures and significant supporting extracellular matrix, 3D cultures are not ideal for host-pathogen studies. Enteroids or colonoids can be grown as epithelial monolayers on permeable tissue culture membranes to allow manipulation of both luminal and basolateral cell surfaces and accompanying fluids. This enhanced luminal surface accessibility facilitates modeling bacterial-host epithelial interactions such as the mucus-degrading ability of enterohemorrhagic E. coli (EHEC) on colonic epithelium. A method for 3D culture fragmentation, monolayer seeding, and transepithelial electrical resistance (TER) measurements to monitor the progress towards confluency and differentiation are described. Colonoid monolayer differentiation yields secreted mucus that can be studied by the immunofluorescence or immunoblotting techniques. More generally, enteroid or colonoid monolayers enable a physiologically-relevant platform to evaluate specific cell populations that may be targeted by pathogenic or commensal microbiota.

Here, we present a protocol to culture human enteroid or colonoid monolayers that have intact barrier function to study host epithelial-microbiota interactions at the cellular and biochemical level.

Human 3-dimensional (3D) enteroid or colonoid cultures derived from crypt base stem cells are currently the most advanced ex vivo model of the intestinal ...

Quantitative Fluorescence In Situ Hybridization (FISH) and Immunofluorescence (IF) of Specific Gene Products in KSHV-Infected Cells

Mechanistic insight arrives from careful study and quantification of specific RNAs and proteins. The relative locations of these biomolecules throughout the cell at specific times can be captured with fluorescence in situ hybridization (FISH) and immunofluorescence (IF). During lytic herpesvirus infection, the virus hijacks the host cell to preferentially express viral genes, causing changes in cell morphology and behavior of biomolecules. Lytic activities are centered in nuclear factories, termed viral replication compartments, which are discernable only with FISH and IF. Here we describe an adaptable protocol of RNA FISH and IF techniques for Kaposi's sarcoma-associated herpesvirus (KSHV)-infected cells, both adherent and in suspension. The method includes steps for the development of specific anti-sense oligonucleotides, double RNA FISH, RNA FISH with IF, and quantitative calculations of fluorescence intensities. This protocol has been successfully applied to multiple cell types, uninfected cells, latent cells, lytic cells, time-courses, and cells treated with inhibitors to analyze the spatiotemporal activities of specific RNAs and proteins from both the human host and KSHV.

Quantitative Fluorescence In Situ Hybridization FISH and Immunofluorescence IF of Specific Gene Products in KSHV-Infected Cells

We describe a protocol utilizing fluorescence in situ hybridization (FISH) to visualize multiple herpesviral RNAs within lytically infected human cells, either in suspension or adherent. This protocol includes quantification of fluorescence producing a nucleocytoplasmic ratio and can be extended for simultaneous visualization of host and viral proteins with immunofluorescence (IF).

Mechanistic insight arrives from careful study and quantification of specific RNAs and proteins. The relative locations of these biomolecules throughout ...

Visualization of Candida albicans in the Murine Gastrointestinal Tract Using Fluorescent In Situ Hybridization

Candida albicans is a fungal component of the gut microbiota in humans and many other mammals. Although C. albicans does not cause symptoms in most colonized hosts, the commensal reservoir does serve as a repository for infectious disease, and the presence of high fungal titers in the gut is associated with inflammatory bowel disease. Here, we describe a method to visualize C. albicans cell morphology and localization in a mouse model of stable gastrointestinal colonization. Colonization is established using a single dose of C. albicans in animals that have been treated with oral antibiotics. Segments of gut tissue are fixed in a manner that preserves the architecture of luminal contents (microorganisms and mucus) as well as the host mucosa. Finally, fluorescent in situ hybridization is performed using probes against fungal rRNA to stain for C. albicans and hyphae. A key advantage of this protocol is that it allows for simultaneous observation of C. albicans cell morphology and its spatial association with host structures during gastrointestinal colonization.

Visualization of Candida albicans in the Murine Gastrointestinal Tract Using Fluorescent In Situ Hybridization

The purpose of this protocol is to visualize Candida albicans cell shape and localization in the mammalian gastrointestinal tract.

Candida albicans is a fungal component of the gut microbiota in humans and many other mammals. Although C. albicans does not cause symptoms in most ...