Is there a difference between ISH and ISHH? (In Situ Hybridization, In Situ Hybridization Histochemistry)

Is there a difference between ISH and ISHH? (In Situ Hybridization, In Situ Hybridization Histochemistry)

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I came across the term ISHH in my document and discovered that it stands for In Situ Hybridization Histochemistry. I's translating to Russian a document that uses this abbreviation.

Example from the PubMed:

We then validated these findings by using a combination of tools, including the analysis of an additional brain region in an independent subject cohort, cross-validation with an alternate custom microarray platform, and follow-up with in situ hybridization histochemistry (ISHH).

Is there any difference between ISH and ISHH, or are these just synonyms for the same method, in situ hybridization?

The term appears to be in use as a synonym. PLOS ONE is a good journal, and in it Okubo, 2016 write "The protocol for ISHH was described in detail previously [22]." Where [22] is Yamanaka, 2007, describing a classic ISH protocol.

Note that both papers use good old fashioned nuclear tract emulsion for S-35 labelled probes. The ISHH protocol is in its own section separate from IHC. It seems a bit odd because no literal "chemistry" is involved in recognizing the target - the RNA probe binds non-covalently and emits radiation. However, as noted in the question discussion, IHC is formally defined (OED) as "the branch of science concerned with the identification and distribution of the chemical constituents of tissues by means of stains, indicators, and microscopy". For example, though I have seen immunofluorescence and immunohistochemistry used to describe mutually exclusive visualization options, noncovalent binding of fluorophore-labelled antibodies is described as "fluorescent IHC".

Current and Emerging Technologies for the Diagnosis of Microbial Infections

Alon Singer , Yi-Wei Tang , in Methods in Microbiology , 2015

4.2 In Situ Hybridisation

In situ hybridisation or ISH is a well-established technique to identify the presence or absence of a particular genomic sequence within a cell. In diagnostic assays, DNA or RNA-based fluorescent probes, are commonly used (termed FISH fluorescent in situ hybridisation) which facilitate cell imaging using a standard fluorescence microscope. As described earlier, the advantage of using artificial NAs such as LNA-modified oligomers and PNAs is the inherent thermal stability of these probes when compared to natural NAs, with the higher thermal stability facilitating the utilisation of shorter probes yielding more specific hybridisation ( Silahtaroglu, Pfundheller, Koshkin, Tommerup, & Kauppinen, 2004 Silahtaroglu, Tommerup, & Vissing, 2003 ). In addition, and unlike natural NA probes, when these probes are to be introduced into live cells, they are highly resistant to enzymatic degradation and can be further modified as needed to increase cellular uptake ( Amann & Fuchs, 2008 ). Unlike microarray technology, there has been an increased interest in developing FISH assays for the use in clinical microbiology particularly targeted to pathogen identification whereas the majority of research work with LNAs has been directed at the detection of human miRNA and mRNA ( Doné & Beltcheva, 2014 Nielsen, 2012 Singh et al., 2014 ) or viral targets ( Cerqueira et al., 2008 Robertson, Verhoeven, Thach, & Chang, 2010 Shiogama et al., 2013 ).

One of the first studies using LNA-modified oligomers for rRNA-targeted FISH assays was described by Kubota and co-workers in 2006 . They described that poor hybridisation efficiency was a significant limitation of FISH methods due to the low affinity of DNA probes ( Kubota, Ohashi, Imachi, & Harada, 2006 ). This study demonstrated that by substituting 2–4 DNA bases within an 18–21mer DNA probe with LNAs, it was possible to produce a signal enhancement ranging from 2 × to over 22 × indicating that simple modifications can result in significant signal enhancement. They also demonstrated that discriminating among targets with two mismatches is feasible with LNA-modified oligomers which is not possible with DNA probes. This corroborates previous studies demonstrating the LNA-modified oligomers outperform natural NAs as FISH probes ( Thomsen, Nielsen, & Jensen, 2005 ). Montone and co-workers further demonstrated this convincingly in formalin-fixed, paraffin-embedded (FFPE) tissue sections by detecting the 18s rRNA of Aspergillus. They demonstrated that standard DNA probes had a significantly weaker signal in the hybridisation reaction than that observed in LNA-modified probes, demonstrating the higher affinity of LNAs to rRNA. Additionally, LNA-modified probes were able to rule out culture-positive cases of Fusarium, where the DNA probes failed as they produced numerous false positives ( Montone, 2009 Montone & Feldman, 2009 ).

Unlike LNA-modified oligomers, PNAs have been utilised more extensively in FISH assays and have made a direct impact in the clinical setting. PNAs for FISH-based assays were introduced shortly after PNAs were first developed, and there have been numerous studies published which describe the optimal PNA oligomer lengths to be between 13 and 18mer, in contrast to DNA probes which range from 20 to 25mer. As a result, PNA-FISH assays display remarkable levels of sequence specificity, potentially at the single-base mismatch level, far superior to that possible with DNA probes ( Cerqueira et al., 2008 ). As the majority of PNA-FISH applications are directed at live cells, PNAs are advantageous as they can be designed to promote cellular uptake (given their relatively hydrophobic nature) and are very resistant to the action of nucleases and proteases. Though not limited to, PNA FISH-based assays have mainly been described for the detection of bacterial pathogens either in FFPE, media, or blood cultures. Many studies have described assays using PNAs for detecting and identifying numerous bacterial and fungal species including of Mycobacterium sp., Staphylococcus sp., E. coli, Pseudomonas sp., Klebsiella sp., Salmonella sp., Listeria sp., Streptococcus sp., Acinetobacter sp., Proteus sp., and others ( Cerqueira et al., 2008 Forrest et al., 2006 González et al., 2004 Lefmann et al., 2006 Montague, Cleary, Martinez, & Procop, 2008 Oliveira, Procop, Wilson, Coull, & Stender, 2002 Oliveira et al., 2003 Peleg et al., 2009 Perry-O'Keefe et al., 2001 Reller, Mallonee, Kwiatkowski, & Merz, 2007 Stender et al., 1999 ). The majority of assays target the rDNA due to its relative abundance, and in the majority of cases, both sensitivity and sensitivity of over 90% were demonstrated. FISH assays using PNAs have been demonstrated to be superior to DNA-based approaches and are likely to be utilised in many more applications in the near future.

Fluorescence In Situ Hybridization (FISH) or Other In Situ Hybridization (ISH) Testing of Uterine Cervical Cells to Predict Precancer and Cancer [Internet].

Cervical cancer screening remains an evolving field with ongoing reevaluation of Pap screening practices and the role of HPV testing, as well as development of new technologies, including ISH testing for genetic abnormalities. The key findings of this review and the strength of evidence are summarized in Table 8.

Table 8

Key Findings and Strength of Evidence.

The horizon scan conducted for Key Question 1 led to the subsequent focus on ISH tests for TERC, MYC, HPV 16, or HPV 18 as tests for cervical abnormalities or cancer.

Our review of data on analytic validity for Key Question 2 revealed a paucity of evidence. We found no studies examining the association between ISH for TERC or MYC and another genetic test in cytology or histology samples. For HPV, we identified some studies for which we could examine the correlation between ISH and reference tests, namely PCR and Hybrid Capture 2. However, these tests measure different biological parameters since, unlike ISH, the reference HPV tests are not restricted to detecting nuclear episomal or integrated HPV. (In situ ISH testing for HPV, which is the only ISH that can identify integration into the genome, may add information beyond the most common ISH testing for 13 or 14 types of HPV or ISH for HPV 16 and 18, which only indicate HPV infection, not integration.)

Further, the panels of HPV genotypes tested for by ISH and the reference tests varied and were not completely overlapping. This heterogeneity limits the conclusions that can be drawn about analytic validity. Not surprisingly, the agreement between ISH tests and reference tests was inconsistent across the studies.

Risk of bias assessment of analytic validity studies showed variable detail of reporting, which was particularly poor for the reference tests. Review of the evidence on thresholds for ISH tests also showed incomplete reporting as well as variable thresholds of positivity and chromosomal control probes used. Information on other preanalytic issues was sparse or not informative. The lack of data on reproducibility is a major deficiency in the evidence base. This suggests a need for research to explore thresholds and standardize test procedures.

For Key Question 3 on clinical validity, the strength of evidence for ISH testing was graded as low, failing to show that the addition of ISH tests resulted in better clinical validity. Clinical practice guidelines suggest that ISH is a potential add-on test after initial Pap testing, with subsequent HPV testing, or after initial Pap and HPV cotesting. In this context, it is more desirable for ISH to show high specificity than high sensitivity. In our review, FISH testing did not show consistently increased sensitivity for the identification of CIN2+ or CIN3+ on histology, although it was more specific than other tests or test combinations. However, we cannot conclude that ISH testing would increase clinical validity of an overall screening strategy. As compared with FISH or Hybrid Capture 2 testing for HPV, FISH for TERC or MYC alone was more specific and less sensitive than the test combinations.

Regarding Key Question 4, we found no studies examining the association of ISH test results with clinical outcomes. There were also no comparative studies of strategies that include ISH tests that examined clinical utility, which would be of particular interest for colposcopy rates and histology results.

Comparison with Current Knowledge

ISH tests are not used routinely used in screening for cervical cancer at this point. However, there is a need to improve the clinical validity of screening for cervical cancer. Thus there is a potential role for tests such as ISH. HPV tests for panels of high-risk genotypes have been shown to have a higher sensitivit, but lower specificity that what we found for ISH tests. 91 Thus when cotesting is used, add-on tests with greater specificity may be useful. However, HPV testing is evolving, and new reference tests for HPV testing will change the performance of add-on tests. ISH may need to be examined as an alternative to tests that can identify HPV 16 and 18 individually. Further, the recent launch of HPV vaccination in adolescents is expected to change the natural history of HPV associated cervical carcinoma going forward.


Formal appraisal of applicability of the Key Question 3 studies on clinical validity with the QUADAS-2 tool showed no major concern regarding applicability. However, studies included populations from around the world, with variable prevalence of HPV infections, CIN classes, and cervical cancer.

CMS has a particular interest in the Medicare population, whose core beneficiaries are 65 years of age or older. On the basis of the lower incidence of HPV infection and cervical cancer among older women who have undergone adequate screening than among younger women, the 2012 guidelines recommend cessation of screening after the age of 65 years (so long as screening tests were negative in the prior 10 years). Since a notable proportion of Medicare beneficiaries are younger than 65, the findings of the report are still relevant for CMS.

Implications for Clinical and Policy Decisionmaking

The current evidence base is insufficient to consider routine ISH testing in the clinical scenarios analyzed in the report. Specifically the evidence is insufficient to recommend routine ISH testing for TERC, MYC, HPV 16 or 18 in women screened or tested for cervical cancer with a finding of LSIL or ASCUS on cytology, with or without HPV infection.


Our review is limited to published reports, which usually do not allow for detailed analysis of individual patient data for subgroups of interest. Studies evaluating more than one test approach did not include cross-tabulation of positive and negative test results across all tests. Our review addresses a limited scope based on what was determined to be the most meaningful clinical questions. Given our stringent inclusion criteria for articles, requiring the mention of cytologic or histologic sampling in the abstract, we may have missed studies that could have contributed additional data for the review of analytic validity.

Regarding Key Question 3 on clinical validity of ISH in particular, the identified evidence base was limited. Studies were generally small and those that we could meta-analyze yielded imprecise effect estimates. Study samples often were from sample banks or databanks, limiting the applicability to the screening population. With one exception, the included studies did not unequivocally report or stratify by HPV status. Studies conducted before the Bethesda terminology change that divided ASCUS into ASC-US and ASC-H may have included a mix of ASC-US and ASC-H in their ASCUS group. There was clinical heterogeneity among the results, given the variety of ISH probe panels used across studies and differences between ISH and the DNA-based reference tests. In addition, the reporting of study quality items was deficient. No studies examined risk prediction with ISH or the test’s clinical utility or addressed screening for cervical adenocarcinoma in particular.

Research Gaps

Our review reveals four major research gaps. First, the assessment of the analytic validity of ISH (Key Question 2) highlights a need to establish common thresholds, probe sets, controls, and procedures. An expert conference may be helpful to agree on common measurement guidelines, a path that was successfully pursued to arrive at the consensus Bethesda classification for cytological abnormalities in cervical cancer. Scoring of ISH slides can be time-consuming. Automated approaches are promising, but in order for ISH to become a routine test, the evaluation of test results needs to be standardized and accelerated.

Second, bigger studies with larger numbers of patients with HSIL are needed to yield more precise estimates.

Third, future research should reflect changes in clinical practice. On the basis of the current guidelines, it can be expected that Pap with reflexive HPV testing or Pap–HPV cotesting will become more widely used. This will require study of the clinical validity of ISH as an add-on test in groups of women characterized as having a normal Pap or ASCUS or LSIL along with a positive or negative HPV test. It is also expected that HPV testing will eventually be able to routinely identify not only high-risk HPV genotypes broadly but also HPV 16 and 18 individually, with the use of either sequential or combined tests. This will require reevaluation of the role of ISH, which we in this review considered as a hypothetical alternative to testing for HPV 16 or HPV 18. Development of automated HPV testing may provide an incentive to explore the performance of up-front HPV testing rather than Pap testing, since testing of cervical cytologic specimens requires a trained human operator. This would generate another constellation in which to study the value of ISH testing added to HPV genotyping.

Fourth, changes in terminology should be followed in future studies, specifically the differentiation between ASC-US and ASC-H and the use of LAST terminology, including p16 staining in ambiguous cases for classifying histology as LSIL or HSIL.

Fifth, further evaluation of clinical validity of ISH should be better designed to achieve this aim. Studies could examine ISH testing for not only a single probe (such as TERC) but also panels of probes, for example for both TERC and HPV. Ideally, large studies would allow for the comparison of multiple tests in order to make it possible to select tests with best analytic validity as well as clinical validity for CIN. However, to measure false negative rates, colposcopy would need to be performed in patients with negative screening tests. Such studies should therefore identify the tests, thresholds, and combinations that are most promising for further evaluation of clinical utility. Efficient exploration of the correct test use (i.e., the testing with the best performance) would again be conducted with several promising tests, thresholds, and test combinations studied simultaneously in a sufficiently large sample on the same specimens and follow patients with routine or test-directed care to assess impacts on diagnostic thinking, evaluation, management, and clinical outcomes. Projecting the clinical utility of different tests may entail modeling of data from different studies in decision analyses.

Lastly, the role ISH testing for detection of adenocarcinoma should be examined. The variability in chromosomal aberrations between squamous-cell cancer and adenocarcinoma suggests that a panel of ISH probes, rather than a single probe, would capture a greater variety of chromosomal changes.


Our report shows an emerging body of literature on ISH testing for cervical cancer. Although ISH tests are marketed by some laboratories for triaging women with abnormal screening tests, there is a lack of standardization of probes and procedures that needs to be addressed. The role of add-on ISH testing has not been adequately examined in current screening contexts, that is, after HPV and Pap testing. Further, HPV testing is likely to evolve, for example with primary screening for 13 or 14 HPV genotypes or with wider availability of HPV16/18 testing. This will again require reexamination of the role of add-on tests such as ISH and its impact not only on diagnostic utility but also on clinical utility (in particular colposcopy) and on clinical outcomes. Thus, the evidence is currently too immature to suggest the ISH testing for routine practice.

Shall you use Biotin or Digoxigenin for ISH Probe Labeling?

ISH, or in situ hybridization, is a powerful technique designed to detect specific nucleic acid sequences within cells and tissue samples. The technique is based on the hybridization between complimentary DNA or RNA probes to a particular target sequence. It is mainly used to detect the presence of specific genomic sequences of any type, including bacteria and viruses. Detection enables the determination of spatial location of sequences within tissues and cells, while also providing insight on relative levels of expression. When stored in optimal conditions, tissue samples can be maintained with minimal signal degradation for years.

The first generation of probes used for ISH were introduced in 1969. These probes were based on radiolabeling technologies, which frequently used isotopes such as 23 P, 35 S, and 33 P. These radioisotopes initially showed promising results displaying high activity and visualization via photographic film exposure. However, these methods of detection were unfortunately hindered by various constraints that reduced practicality and convenience. Isotopic probes are inherently unstable due to the relatively short half-lives of their associated isotope labels. This means that tissue samples are usually short lived and cannot be maintained for extended periods of time. Even more concerning is the potential hazards implicated on end users due to radioactive exposure, necessitating more stringent protection.

Because of these potential hazards and drawbacks, developments were made to create alternative options such as non-isotopic probes using other haptens. Two common non-isotopic haptens used to generate ISH probes are biotin and digoxigenin. Detection mechanisms typically utilize fluorescence or enzymatic colorimetric development through antigen-antibody interaction in the case of digoxigenin and streptavidin affinity binding to biotin. Methods of creating non-isotopic probes usually require incorporation of conjugated UTP into sequences. This can be conducted through nick translation or random priming. While direct conjugation of probes using Alkaline phosphatase (AP) or fluorophores is possible and has existed, it is not used as frequently due to expensive costs. Overall, non-isotopic haptens are known to have better resolution, stability, and shorter development times compared to their isotopic counterparts.

The first non-isotopic hapten to find widespread success were biotin-conjugated ISH probes. Biotinylated probes are further enhanced by higher affinity binding to streptavidin, which is usually conjugated with alkaline phosphatase for color development. Biotinylated probes are usually easier to work with and are less expensive to generate in contrast to isotopic variants. Because radioactive decay is not a factor, biotinylated probes can be retained in tissue samples for extended periods of time as long as they are stored in appropriate conditions protected from light. Despite this, there are several known inefficiencies unique to biotinylated systems. Traditionally, biotin could be incorporated through direct in-vitro transcriptional reactions via biotin-coupled rNTPs. Alternatively, incorporation can also be made through allylamino-UTP with subsequent coupling of the allylamino group to biotin-N-hydroxysuccinimide. However, both processes have shown notable differences in transcription rates due to steric hindrances of conjugated molecules, resulting in suboptimal nucleotide interaction with transcriptional machinery. It is also known that tissue samples contain endogenous levels of biotin. Since the conjugated streptavidin associates with biotin at a high affinity, the possibility for producing high background and false positives exists.

Figure 1: Workflow of hybridization with DIGX® HPV Probes and detection with DIGX® linkers and POLYVIEW® PLUS reagents.

Figure 2: DIGX® HPV type 16/18/31/33/51 probe (ENZ-GEN114) was used to assay cervical tissue using a Leica Bond III. Image was acquired with 10x objective.

Materials and Methods

Batrachochytrium dendrobatidis (Bd) and B. salamandrivorans (Bsal) Culture

Bd isolate ALKL1 (isolated from an eastern newt [Notophthalmus viridescens] from Virginia, USA) and Bsal isolate AMFP (isolated from a morbid wild fire salamander [Salamandra salamandra] from Bunderbos, The Netherlands) were grown on 1% tryptone plates (1, 23). The cultures were individually scraped and placed in a 1.5 ml microcentrifuge tube containing approximately 0.7 ml 70% ethanol. Additional aliquots of scraped Bd and Bsal were combined and fixed in 70% ethanol together. Microcentrifuge tubes were briefly centrifuged, ethanol was removed, and the fixed fungi were suspended in Histogel (Richard-Allan Scientific, Kalamazoo, Michigan), processed routinely and embedded in paraffin.

Experimental Infection of Eastern Newts (Notophthalmus viridescens) With Bd and Bsal

Adult eastern newts were collected from Maryland, screened for Bd and Bsal, treated with itraconazole, and experimentally infected with Bd, Bsal, or both Bd and Bsal as previously described (23). All Batrachochytrium infected newts died during the experiment, and uninfected control newts were euthanized using benzocaine hydrochloride. Postmortem swabs were screened for Bd and Bsal by qPCR as previously described (23). Newts were collected under MD Department of Natural Resources permit No. 56,427. Lab protocols were performed under University of Maryland IACUC protocol R-15-15.

Newts were fixed in 10% neutral buffered formalin for 繈 h before being transferred to 70% ethanol. Two newts from each experimental group (Bd-/Bsal- Bd+/Bsal- Bd-/Bsal+ Bd+/Bsal+) were decalcified in 0.5 M ethylenediamine tetraacetate acid (EDTA), ph 8.0 for 縤 h. Serial transverse sections of the body and sagittal sections of all limbs and the tail were taken and submitted for routine processing and paraffin embedding.

Experimental Infection of Large-Blotched Ensatinas (Ensatina eschscholtzii klauberi), Red Salamanders (Pseudotriton ruber), Eastern Newts (Notophthalmus viridescens), Red-Legged False Brook Salamanders (Aquiloeurycea cephalica), and Yellow-Eyed Ensatinas (Ensatina eschscholtzii xanthoptica) With Bsal

Large-blotched ensatinas (E. e. klauberi) and red salamanders (Pseudotriton ruber) were raised in captivity by Indoor Ecosystems LLC (Whitehouse, OH, USA). Eastern newts (N. viridescens), yellow-eyed ensatinas (Ensatina eschscholtzii xanthoptica), and red-legged false brook salamanders (Aquiloeurycea cephalica) and were collected from the wild (Tennessee, USA, California, USA, and Mexico, respectively). Notophthalmus viridescens were collected under Tennessee Wildlife Resources Agency Scientific Collection Permit �. Ensatina eschscholtzii xanthoptica were collected under California Department of Fish and Wildlife Scientific Collection Permit #SC-11505. Aquiloeurycea cephalica were collected under permit from the Agriculture, Stockbreeding, Rural Development, Fishing and Food Ministry of Mexico and import permit #MA87825B-1 from the United States Fish and Wildlife Service. The A. cephalica were determined to be naturally infected with Bd by qPCR using previously described methods, which provided an opportunity for a natural co-exposure experiment (24). All other animals were verified as Bd qPCR negative.

Bsal isolate AMFP was grown on tryptone gelatin hydrolysate plates and flooded to create serial dilutions from 5 × 10 3 to 5 × 10 6 Bsal zoospores/mL. Individuals from each species were exposed to Bsal in 100-mL containers for 24 h that contained 1 mL of inoculum and 9 mL of autoclaved water. After the Bsal challenge, salamanders were housed in husbandry containers with moist paper towel and a cover object. Salamanders were humanely euthanized using benzocaine hydrochloride upon loss of righting ability. All husbandry and euthanasia procedures followed the Amphibian Husbandry Resource Guide of the Association of Zoos and Aquariums and the Guide for Euthanasia by the American Veterinary Medical Association. All animal procedures were approved by University of Tennessee Institutional Animal Care and Use Committee protocol �.

At necropsy, all animals were swabbed to test for the presence of Bsal DNA (and Bd for A. cephalica co-exposure experiments). Genomic DNA was extracted (Qiagen DNeasy Blood and Tissue Kit, Hilden, Germany) and quantitative PCR performed using an Applied Biosystems Quantstudio 6 Flex system (Thermo Fisher Scientific, USA) using previously described methods (24). All swabs were run in duplicate, and the number of Batrachochytrium zoospore copies/μl was calculated using serial dilutions of synthetic Bsal and Bd DNA.

After necropsy, samples were preserved in 10% neutral buffered formalin until processing (2� days). The length of formalin storage was varied to examine potential effects of fixation time on ISH sensitivity. Animals were then decalcified in 0.5M EDTA, ph 8.0 for 縤 h. Serial transverse sections of the body and sagittal sections of all limbs and the tail were taken and submitted for routine processing and paraffin embedding.

Naturally Occurring Little Devil Poison Frog (Oophaga sylvatica) Chytridiomycosis

A captive born and bred little devil poison frog (Oophaga sylvatica) was submitted to the Aquatic, Amphibian, and Reptile Pathology Service at the University of Florida's College of Veterinary Medicine for diagnostic necropsy. The frog arrived in 70% ethanol, was transferred to 10% neutral buffered formalin for 24 h, and then decalcified in 0.5M EDTA, ph 8.0 for 縤 h. Serial transverse sections of the body and sagittal sections of all limbs and the tail were taken and submitted for routine processing and paraffin embedding. Histopathologic findings were consistent with Bd chytridiomycosis, and the sample was utilized for ISH as an anuran (frog) chytridiomycosis sample.

RNAScope ® Dual-Plex Chromogenic in situ Hybridization (ISH)

Custom RNAScope ® target-specific oligonucleotide (ZZ) probes were designed by Advanced Cell Diagnostics (Advanced Cell Diagnostics, Newark, California). Two ZZ probes, complementary to nucleotides 452-555 of B. dendrobatidis JEL 197 (NG_027619.1) and nucleotides 444-584 of Batrachochytrium salamandrivorans isolate AMFP (KC762293.1) 28S rRNA sequences, were designed each for Bd and Bsal, respectively. Chromogenic ISH was performed on a Leica BOND RX Fully Automated Research Stainer (Leica Biosystems, Buffalo Grove, Illinois) using RNAscope technology (22). Formalin-fixed paraffin-embedded tissue blocks were sectioned at 4 μm and mounted on Fisherbrand SuperFrost Plus glass slides (Fisher Scientific, Pittsburgh, PA). Automated, dual-plex, chromogenic RNAscope using methodology based on Anderson et al. (20) was performed using both Bd [LS, catalog �] and Bsal [LS, catalog �-C2] ISH probes. Pretreatment, hybridization, signal amplification, and detection (RNAscope Reference Guide, ACD/Biotechne) were performed on the Leica BOND RX. Pretreatment conditions included sequential deparaffinization, target retrieval using Leica Epitope Retrieval Buffer at 95ଌ for 15 m, protease digestion at 40ଌ for 15 min, and endogenous enzyme block. Bd and Bsal probes were hybridized at 42ଌ for 120 min, followed by signal development. After hybridization, serial signal amplification reactions were followed by online fast red chromogenic development of the Bsal probe. PermaGreen/HRP (Diagnostic Biosystems, Pleasanton, CA) was manually applied for 3 min to visualize the Bd probe. Slides were counterstained with hematoxylin, ImmunoHisto-Sealer (Diagnostic BioSystems, Pleasonton, CA) was applied for long-term retention of the green signal, and coverslips were mounted with Vectamount Permanent (Vector Laboratories, Burlingame, CA). The dihydrodipicolinate reductase (dapB) probe [LS, catalog �] was used as a negative control probe and was applied to separate histologic sections processed concurrently and in the same manner as those that received the Bd and Bsal probes. Glass slides were visualized on an Olympus BX43 microscope (Olympus Corporation, Tokyo, Japan) photomicrographs were captured with a Spot Insight 12 Mp sCMOS Color Camera (Spot Imaging, Sterling Heights, Michigan) and SPOT Imaging software (v5.4.3 Spot Imaging, Sterling Heights, Michigan).

Introduction to RNA FISH

Fluorescent in situ hybridization targeting ribonucleic acid molecules (RNA FISH) is a methodology for detecting and localizing particular RNA molecules in fixed cells. This detection utilizes nucleic acid probes that are complementary to target RNA sequences within the cell. These probes then hybridize to their targets via standard Watson-Crick base pairing, after which one may detect them via fluorescence microscopy, either through direct conjugation of fluorescent molecules to the probe, or through fluorescent signal amplification schemes. Recent advances in RNA FISH have increased the specificity and sensitivity of the method to enable the detection of individual RNA molecules, providing very accurate measurements of of RNA abundance and localization at the single cell or even subcellular level. While most applications thus far have been in fixed cells, advances in probe technology have lead to the ability to detect single RNA molecules in living cells.

The protocols for in situ hybridization (ISH) targeting RNA molecules are well established and conceptually simple. All RNA ISH protocols essentially involve bathing the sample in a high concentration of nucleic acid probe (or probes) that are complementary in some way to the target RNA species, driving hybridization of the probe to the target, a principle derived from the development of DNA ISH (Gall and Pardue, 1969). After hybridization, one washes away excess unbound probe, theoretically leaving only those probes specifically bound to the target molecule. Differences between the variants of RNA ISH typically revolve around the type of nucleic acid used for the probe and the type of labeling scheme used to detect the probe via microscopy. Initially, researchers used radiolabeled cDNA probes complementary to the appropriate target (Harrison et al., 1973). Issues with radiolabeling include the low spatial resolution and difficulties associated with handling and stability of radioactive materials. Thus, the development of fluorescence-based (FISH) approaches using DNA or RNA probes provided a major step forward in the field, first applied to DNA FISH (Bauman et al., 1980) and then RNA FISH (Singer and Ward, 1982). Rapidly, researchers adopted the approach of generating cDNA or RNA probes (via enzymatic amplification or nick translation) containing modified bases (Langer et al., 1981) that allowed the conjugation of various haptens or even fluorophores, thus facilitating either indirect or direct detection via fluorescence microscopy. These probes did present some challenges, however, because it was hard to ensure a consistent and high degree of labeling from experiment to experiment. Moreover, the sensitivity of that style of fluorescent probes is generally poorer than that of radiolabeled probes due to cellular autofluorescence. This is problematic because many important mRNAs (such as those encoding transcription factors) are often present at very low abundances, often on the order of a few molecules per cell or less. Subsequent developments in RNA FISH methodology, however, have resolved many of these problems by refining RNA FISH to the point where it can detect single RNA molecules, enabling the direct quantification of RNA species of very low abundance and providing absolute measurements of RNA copy number.

Broadly, current methods for single molecule RNA FISH fall into two categories: those that use some form of signal amplification, and those that rely on direct detection of signal. Direct detection involves labeling the probes themselves with fluorophores. In order to achieve single molecule sensitivity, the probes must have enough fluorescence to be detectable above background autofluorescence. One technique is to use a set of short single-stranded DNA oligonucleotides complementary to various regions of the target RNA, each labeled with one or more fluorescent moieties (Femino et al., 1998 Raj et al., 2008). The binding of multiple probes localizes enough fluorophores to the target RNA such that the RNA is easily visible as a fluorescent spot via fluorescence microscopy (Fig. 1). The benefit of using several oligonucleotide probes at the same time is that the off-target binding of a single oligonucleotide in the probe pool will either be undetectable or readily distinguishable to the much brighter spots corresponding to the true RNA, thus reducing the chances of false positives. False negatives are similarly unlikely, for even if a single probe out of the pool fails to bind, the rest are likely to bind. Recent research indicates that the use of nucleic acid chemistries with tighter and more specific binding properties such as linked nucleic acids (LNA) may be able to produce similar results using a single probe (Taniguchi et al., 2010), although the detection of just one or few fluorescent molecules may only be feasible in cases like bacteria where cellular autofluorescence is kept to a minimum (as opposed to probe pools, which work in a variety of contexts (Raj et al., 2008)). In general, the signals produced via these methods are low and require the use of sensitive CCD cameras and high NA optics for detection.

In order to circumvent the limitation of low signals from the relatively small numbers of fluorescent molecules targeted to mRNA in these direct detection methods, researchers have also developed a large variety of schemes to amplify signals from individual molecules. Some of these are relatively simple extensions of the direct detection methods, such as detection of the probe by fluorescently labeled antibodies targeting specific haptens incorporated in large numbers into an RNA probe (Paré et al., 2009). Others involve targeting nucleic acid probes with a single or few haptens with antibodies conjugated to enzymes those enzymes in turn will act upon a substrate in such a manner as to create a fluorescent product that will become covalently linked to surrounding molecules (Kerstens et al., 1995). Yet other methods amplify signals by using DNA polymerase and circular templates to locally create long, repetitive single-stranded DNA tracts in situ, which one then targets with short oligonucleotide probes (Larsson et al., 2010). Such methods have the advantage of labeling targets to such a degree that the signals are easily visible even by eye, precluding the need for expensive optical setups. Also, they are able to reliably detect short RNA molecules such as miRNA (Lu and Tsourkas, 2009), and researchers have even demonstrated the ability to detect single-base differences in RNA molecules (Larsson et al., 2010). However, such methods are somewhat prone to lower detection efficiencies owing to the large number of steps in such protocols, each of which has some probability of failure. Some reports indicate that such issues are relatively minor (Lu and Tsourkas, 2009), while others amplification methods detect only a small fraction of the target mRNA (Larsson et al., 2010). Another point of comparison between direct detection and amplification is the ability to detect different species at the same time by using spectrally distinguishable fluorescent moieties: many amplification methods are limited to a single “color” owing to the necessity of using a single enzyme/substrate pair, whereas direct detection can utilize the plethora of organic dyes with different spectral properties currently available. The adoption of these various methods for performing FISH in fixed cells are likely to depend on the particular demands of the application at hand.

Traditionally, researchers have considered FISH to be a methodology that only applies to fixed cells, as the use of oligonucleotides in living cells has proven to be very challenging for a host of reasons. Chief among these are the fact that cells quickly sequester short single stranded DNA oligonucleotides in the nucleus for rapid degradation, and the inability to wash away unbound probe, leading to high background. A recent study has circumvented these issues by designing a degradation-resistant variant of a “molecular beacon”, which is a short oligonucleotide probe with a fluorophore on one end and a quencher on the other (Tyagi and Kramer, 1996). The probe is designed in such a way that the fluorophore and the quencher are in close proximity when the probe is not bound to the target but are far apart when it is bound to the target thus, the probe is only fluorescent when bound to the target. This reduces the background to the point where one can detect individual mRNA molecules in living cells (Vargas et al., 2005). Further refinements of these methods may open the door to new applications of these live cell FISH techniques.

RNA FISH has a large number of biological applications, but is particularly useful as an assay for spatial aspects of gene expression. One scenario is one in which there is significant cellular heterogeneity, precluding more traditional bulk biochemical assays such as qRT-PCR or northern blots. For instance, it is commonly used to detect gene expression in the study of developmental biology, where it allows for the detection of gene expression in particular sets of cells within the context of a whole developing organism. It also has applications in fully developed tissue that contains a variety of cellular types that would otherwise be averaged together. It is also useful in the study of subcellular localization of RNAs. Examples include the asymmetries in mRNA distributions in the developing Drosophila melanogaster syncytium (Fowlkes et al., 2008), the localization of ASH1 mRNA to the daughter cell during yeast cell division (Long et al., 1995), and the localization of beta-actin mRNA to the leading edge of cells during cellular migration (Lawrence and Singer, 1986). The development of single molecule RNA detection via FISH allows for the accurate quantification in these contexts, and allows for the detection of the localization of even low abundance mRNAs and non-coding RNAs (Khalil et al., 2009). Single molecule counting also provides absolute quantification, which is also useful even in cases like cell lines in which bulk methods are in common use, providing, for instance, very accurate measurements of variability in gene expression (Raj et al., 2006 Raj and van Oudenaarden, 2009). On the horizon, the development of live single-molecule RNA FISH methods promise to bring a much deeper temporal understanding of transcription and RNA dynamics.

The advent of single molecule sensitivity represents in many ways the ultimate fruition of RNA FISH. Through the establishment of simple, reliable methods, RNA FISH represents in many ways a new gold standard for RNA quantification. On the horizon, the development of live cell versions of FISH promise to animate the already rich three-dimensional picture RNA FISH can yield in fixed cells. With applications across a range of biological fields, RNA FISH is a powerful tool for studying gene expression and RNA biology.

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Results and discussion

Formamide bleaching increases signal intensity

Achieving maximal signal intensity in WISH requires balancing preservation of target mRNA with permeabilization of tissue to allow probe hybridization. Using the planarian WISH protocol established in [21] as a starting point, we began systematically testing modifications to improve signal sensitivity with the goal of improving detection of problematic transcripts by FISH. Because the TSA reaction used for fluorescent detection of transcripts rapidly proceeds to completion, we began by using alkaline phosphatase-based detection to directly compare the rate of development of various probes while varying conditions including fixation, bleaching, permeabilization, hybridization buffer, and hybridization temperature. We first tested the effects of these variations using readily detected transcripts, including the neoblast marker smedwi-1[24] moderately detected transcripts, including a vacuolar ATPase B subunit we have identified as being upregulated in the intestine (Smed-vatpaseB), and the midline marker Smed-slit-1[25] and with weakly detected transcripts including the hunchback-like transcription factor, Smed-hb, reported to be broadly expressed [15].

Most of the variations we tested had minimal impact on signal intensity. However, we found that replacing the overnight peroxide bleach in methanol with a short peroxide bleaching step in formamide dramatically reduced development time for all probes tested, indicating improved signal sensitivity (Figure 1A-P). For FISH, the increased signal intensity resulting from the short formamide bleach also improved the signal-to-noise ratio (Figure 1Q and R). Additionally, planarians bleached in formamide showed more consistent labeling of the prepharyngeal region, a densely packed area with typically reduced probe penetration, compared to methanol-bleached planarians, suggesting that tissue permeability was improved. We examined whether a reduction step [21], which was added to improve permeability of the prepharyngeal region, was required in formamide-bleached planarians. Surprisingly, we found that the reduction step slightly diminished signal intensity (Figure 1A-P). If peroxide bleaching in formamide functions to improve tissue permeability, signal intensity should gradually increase with longer bleaching times and eventually reach a maximum level of signal. Consistent with this, we noticed signal intensity improved dramatically after bleaching for 30 minutes, reaching a maximum between 1 to 2 hours incubation in formamide bleaching solution (Figure 1S, U, and W). Interestingly, the improved signal intensity resulting from bleaching in formamide is lost when animals are first bleached overnight in methanol (Figure 1T, V, and X). One possibility for this could be damage of target mRNAs during the long methanol bleaching step. However, when we compared unbleached animals to animals bleached overnight in methanol we observed similar signal intensity (not shown). Additionally, while planarians bleached overnight in formamide had slightly more diffuse signal, signal intensity was similar to animals bleached for two hours in formamide (not shown). These results suggest mRNA is relatively stable during an overnight peroxide bleach, and that pre-bleaching in methanol must mask the benefit of bleaching in formamide through some other mechanism. Interestingly, signal intensity for WISH increases slightly in zebrafish following peroxide bleaching in methanol [26]. Our finding of enhanced signal intensity with formamide-bleaching could be directly and broadly beneficial in improving WISH signal in other organisms, whether they are pigmented or not.

Bleaching animals in formamide improves WISH and FISH signal. (A-P) Chromogenic detection of smedwi-1 (A-D), Smed-vatpaseB (E-H), Smed-slit-1 (I-L), and Smed-hb (M-P) in planarians fixed with or without a reduction step and bleached either overnight in methanol or for 2 hours in formamide as indicated. Development for each probe was stopped at the same time to allow for direct comparison in signal intensity between conditions. (Q-R) Single confocal sections showing FISH for smedwi-1 in the prepharyngeal region of planarians bleached overnight in methanol (Q) or for 2 hours in formamide (R). (S-X) Maximum intensity projections of whole planarians with FISH for Smed-CAVII-1. Planarians were either bleached in formamide alone for 30 minutes (S), 1 hour (U), or 2 hours (W), or pre-bleached overnight in methanol and then bleached again for 30 minutes (T), 1 hour (V), or 2 hours (X) in formamide. Confocal images were false colored in ImageJ using the fire look up table where blue = weak signal and white = strong signal. Scale bars: 500 μm (A-P and S-X) 100 μm (Q-R).

Modified blocking and wash buffers dramatically improve signal specificity

One of the challenges in achieving high signal sensitivity for FISH is that the TSA reaction proceeds rapidly to completion and cannot be monitored and stopped when an optimal signal-to-noise ratio has been reached. Therefore, eliminating weak background staining is vital for optimal signal sensitivity when using the TSA reaction for FISH.

To further improve signal sensitivity we next examined different blocking and wash solutions. A variety of different blocking and wash solutions have been employed for FISH in other systems [21, 26–28]. We began by comparing the effect of adding various reagents to the blocking buffer. Since different antibodies can respond differently to changes in blocking solution, we tested modified blocking solutions with anti-digoxigenin- (DIG), anti-dinitrophenol- (DNP), and anti-fluorescein- (FAM) antibodies conjugated to peroxidase. We found that addition of either casein or PerkinElmer Blocking Reagent (PEBR) improved the signal-to-noise ratio for most of the antibodies tested, but also led to slightly reduced signal intensity (Figure 2). Impressively, addition of Roche Western Blocking Reagent (RWBR) dramatically reduced background, particularly for the anti-DIG and anti-FAM antibodies, without significantly affecting signal intensity (Figure 2).

Addition of blocking reagents greatly improves signal specificity for peroxidase-conjugated anti-hapten antibodies. Planarians were hybridized with DIG-, DNP-, or FAM-labeled anti-sense RNA probes to Smed-histoneH2B (as indicated) and then blocked in buffer with either 5% horse serum alone (A, B, and C), or in 5% horse serum with 0.75% casein (D, E, and F), 0.75% PerkinElmer Blocking reagent (PEBR) (G, H, and I), or 0.75% Roche Western Blocking reagent (RWBR) (J, K, and L) prior to incubation with the appropriate peroxidase-conjugated anti-hapten antibody in the same blocking buffer. Images are maximum intensity projections of planarian heads imaged using identical settings and false colored in ImageJ using the fire look up table. Scale bar: 100 μm.

Blocking and wash solutions for whole-mount FISH in a variety of animals differ in the use of maleic acid, phosphate, or Tris as the buffering component, but typically contain Tween 20 as a detergent [21, 26–28]. We did not observe significant differences between the use of different buffering reagents, simplifying the protocol by reducing the number of stock solutions to prepare and allowing for the use of solutions that are more convenient to make or are already at hand. Significantly, we did find that altering the detergents present in the blocking and wash solutions further improved signal specificity. Addition of, or substitution with, 0.3% Triton X-100 resulted in a slight but noticeable improvement in signal (Figure 3). The benefit was especially pronounced with the anti-DIG and anti-FAM antibodies. There are relatively few peroxidase-conjugated anti-hapten (e.g. DIG, DNP, and FAM) antibodies available that are suitable for TSA, and the reagents described here are widely used for FISH in other model systems. While every model system presents its own unique requirements, the modified blocking solution we present here should have wide utility for the use of these anti-hapten antibodies in other organisms. Chromogenic WISH using alkaline phosphatase-based reagents has minimal background staining compared to FISH, and not surprisingly we observed little difference when RWBR or Triton X-100 was used for chromogenic detection (not shown).

Addition of Triton X-100 to blocking and wash buffers improves signal-to-noise ratio for peroxidase-conjugated anti-hapten antibodies. Planarians were incubated, as indicated, with DIG-, DNP-, or FAM-labeled anti-sense RNA probes to Smed-PKD1 and then blocked in 5% horse serum with 0.5% RWBR, incubated in anti-hapten antibody, and washed in Tris-sodium chloride buffer containing either 0.05% Tween (A, B, and C), 0.3% Triton X-100 (D, E, and F), or 0.05% Tween and 0.3% Triton (G, H, and I). Images are maximum intensity projections of planarian heads imaged using identical settings and false colored in ImageJ using the fire look up table. Scale bar: 100 μm.

Quenching endogenous autofluorescence with copper sulfate

Planarians exhibit autofluorescence over a broad range of wavelengths, and this feature has been used to distinguish newly regenerated tissues from more mature tissues [14]. When we compared the intensity of autofluorescence at various wavelengths between planarians incubated overnight in hybridization buffer at room temperature (Figure 4A, D, G, and J) or at 56°C (Figure 4B, E, H, and K) we noticed an increase in autofluorescence at several wavelength ranges in animals incubated at 56°C. As higher levels of autofluorescence reduce the signal-to-noise ratio, it can be difficult to distinguish real FISH signal from background autofluorescence, especially for low-abundance transcripts. One approach for improving the signal-to-noise ratio for FISH experiments is to use longer wavelength fluorophores for weakly detected transcripts, as autofluorescence tends to be stronger in the blue-to-green range of the spectrum.

Autofluorescence is reduced following treatment with copper sulfate. (A-L) Single confocal sections of the prepharyngeal region of planarians excited with 405 nm (A, B, and C), 488 nm (D, E, and F), 561 nm (G, H, and I), and 633 nm (J, K, and L) laser lines. Planarians exhibit significant autofluorescence at several wavelengths (A, D, G, and J) that increases following a 16 hour incubation at 56°C (B, E, H, and K). Increased autofluorescence is significantly reduced at all wavelengths following a 1 hour incubation in copper sulfate solution (C, F, I, and L). (M and N) Maximum intensity projection of the dorsal posterior of a planarian following FISH for Smed-EGFR-5 before (M) or after (N) copper sulfate quenching of autofluorescence. Arrows indicate the same position in M and in N. Scale bar: 100 μm.

An additional approach for improving signal sensitivity is to reduce or eliminate autofluorescence. While there are a variety of causes for autofluorescence, the broad range of autofluorescence in planarians, its increase following incubation at high temperatures, and its resistance to photobleaching (not shown) is similar to lipofuscin-based fluorescence observed in tissues of other animals [29, 30]. Incubation in copper sulfate solution has been reported to quench lipofuscin-based autofluorescence [29, 30]. To test the ability of copper sulfate to reduce background signal in planarians, we incubated heat-treated animals for 1 hour in copper sulfate solution (10 mM CuSO4, 50 mM ammonium acetate pH 5.0) and imaged using identical settings to the unheated and heat-treated samples. The copper sulfate treatment dramatically reduced autofluorescence at all wavelength ranges examined (Figure 4C, F, I, and L).

The nearly complete elimination of autofluorescence we observed was very encouraging. However, treatment with copper sulfate has been reported to quench some fluorophores [29]. To test whether the benefits of copper sulfate outweigh its potential harm to signal, we analyzed expression of Smed-EGFR-5 (EGFR-5), which is detected at moderate levels in protonephridia [14], using TAMRA-conjugated tyramide before and after copper sulfate treatment. Prior to quenching, detection of EGFR-5 in protonephridia was discernible, but autofluorescence in the secretory cells, which have a similar tubular pattern, complicated visualization of signal (Figure 4M). When we imaged the same animal after treatment with copper sulfate we had to increase the gain to achieve a similar level of brightness. However, the signal-to-noise ratio was dramatically improved, greatly facilitating visualization of EGFR-5 (Figure 4N). The significantly enhanced signal-to-noise ratio we observed for EGFR-5 highlights the utility of copper sulfate treatment for analyzing the expression pattern of transcripts with weak-to-moderate signals. Copper sulfate treatment should also be useful for multicolor fluorescence experiments, as we have noticed only minor quenching of DyLight 405-, FAM-, Cy3-, and DyLight 633-tyramides as well as Alexa488-conjugated secondary antibodies following copper sulfate treatment (not shown).

Balancing signal sensitivity while preserving tissue morphology in regenerates

FISH analysis of planarians within the first few days following amputation presents a challenge, as the blastema tissue is particularly fragile and can be easily damaged by the aggressive treatments required to permeabilize mature tissues sufficiently. Therefore, it is important to achieve a balance during the permeabilization steps that allows for relatively even penetration of probe into mature tissues without excessively damaging blastema tissue. One strategy for accomplishing this is to adjust Proteinase K concentration and incubation time until a satisfactory result is obtained. Additionally, while experimenting with alternative methods for permeabilizing planarians, we noticed that heat-induced antigen retrieval (HIAR) resulted in slightly weaker signal in intact planarians, but allowed for consistent and even labeling throughout the animal while causing less damage to superficial layers compared to Proteinase K treatment (not shown).

We decided to see whether HIAR would achieve the desired balance between permeabilization and preservation of tissue morphology in regenerates. For this purpose we performed FISH on planarians fixed three days after amputation using the neoblast marker smedwi-1[24] and a differentiation marker, Smed-AGAT-1 (AGAT-1) [31], which is expressed in superficial cells just basal to the epidermis. Planarians were processed in parallel and were either permeabilized with Proteinase K treatment followed by post-fixation, or were boiled in sodium citrate buffer for 10 minutes and then incubated in Phosphate Buffered Saline [32] containing 0.3% Triton X-100 and 1% SDS for 20 minutes at room temperature. Single confocal sections of central focal planes reveal strong and even labeling of neoblasts with smedwi-1 in intact tissues for both treatments (Figure 5A and B, magenta). However, blastema morphology was better preserved in planarians treated by HIAR, as the epidermis and superficial layer of AGAT-1-expressing cells are retained, and density of nuclei is higher. We also observed less damage to blastema tissue in planarians treated by HIAR in chromogenic WISH (not shown). In addition to the benefit of HIAR for FISH of regenerating planarians, this method may also be useful for immunostaining following FISH as Proteinase K treatment can destroy epitopes for some antibodies.

Improved signal intensity in regenerates and for low-abundance transcripts . (A-B) Single confocal sections of three day regenerating planarians labeled with DAPI (blue), the neoblast marker smedwi-1 (magenta), and the differentiating cell marker Smed-AGAT-1 (green). While gene expression is detectable in the intact tissue of planarians permeabilized with Proteinase K (A) or following HIAR (B), the superficial layers of the blastema (asterisk), including the AGAT-1-expressing cells, are lost following Proteinase K treatment. Dashed line denotes plane of amputation. (C-D) Maximum intensity projection of animals hybridized with DIG-labeled RNA probes for Smed-nog1 with a standard single TSA using TAMRA-conjugated tyramide (C) or iterative TSA first using DNP-tyramide followed by washing, incubation with peroxidase-conjugated anti-DNP antibody, and a second TSA reaction with TAMRA-conjugated tyramide (D). (C’-D”’) Maximum intensity projections of regions from the planarians in C and D, including the brain (C’ and D’), ventral nerve cord (C” and D”) and the lateral margin (C”’ and D”’). Scale bars: 100 μm.

Enhancing signal intensity through iterative TSA

For particularly low-abundance transcripts, gene expression patterns can be especially difficult to determine due to low signal intensity. While new and more sensitive imaging systems are vastly improving the ability to image weak signals, it can still be difficult to rapidly screen expression patterns in multiple samples by epifluorescence to identify animals or regions on which to focus imaging efforts. For example, Smed-nog1 (nog1) is weakly to moderately detected in portions of the central nervous system, around the body margins, at the base of the pharynx, and at the mouth [23, 33]. nog-1 FISH signal following a conventional single TSA is challenging to discern when viewed by eye under epifluorescence, but yet is capable of being detected by confocal microscopy (Figure 5C). In an attempt to boost signal intensity for nog1 we performed TSA first with DNP-conjugated tyramide, then incubated with peroxidase-conjugated anti-DNP antibody followed by a second, iterative, TSA with fluorophore-conjugated tyramide. In the first reaction signal is amplified by covalently depositing multiple DNP-tyramide molecules near the site of antibody binding. The signal is then further amplified by localizing additional peroxidase-conjugated antibody to the sites of DNP deposition and then performing an additional amplification with fluorophore-conjugated tyramide, which can then be visualized. When we performed iterative TSA for nog1 we noticed a dramatic increase in signal intensity that greatly facilitated observation (Figure 5D). At higher magnification it is easy to identify nog1-positive cells near the cephalic ganglia in animals processed with iterative TSA (Figure 5D’), whereas with single TSA, signal is detected just above background (Figure 5C’). nog1 detection is stronger in the ventral nerve cords, and easily observed following either single or iterative TSA (Figure 5C” and D”). The weak expression in cells around the body margin is almost undetectable following single TSA (Figure 5C”’), but after iterative TSA, nog1-expressing cells are easily identified (Figure 5D”’). The improved specificity from the optimized blocking and wash buffers has been particularly beneficial to iterative TSA, as minor background from non-specific antibody binding is greatly amplified with this method. While the extensive washing following TSA with DNP-conjugated tyramide appears critical, we have had success deploying this technique in multicolor FISH experiments without greatly extending the length of the experiment (see Additional file 1 for details).

Azide effectively quenches peroxidase activity without inhibiting subsequent gene detection in multicolor FISH

Besides providing excellent spatial resolution of gene expression, FISH has particular utility in determining the expression of genes relative to one another. For multicolor FISH to be effective, peroxidase activity of the first antibody used must be quenched effectively prior to subsequent detection rounds without leading to progressive degradation of sample. A number of methods have been described for inactivating peroxidase activity in multicolor FISH experiments, including incubation with hydrogen peroxide [21], fixation with formaldehyde [34], incubation in low pH buffer [35], and treatment with azide [36]. While incubation with hydrogen peroxide is more widely used, there does not seem to be a consensus on which method is the most effective. Therefore, we decided to compare directly several methods to determine which was the most effective at inactivating peroxidase activity and least detrimental to detection of subsequent gene expression patterns in planarians. For this we performed multicolor FISH for two non-overlapping genes Smed-CAVII-1 (CAVII-1), which is detected at high levels in the protonephridial system [37], and a homolog of Polycystin 1, Smed-PKD1 (PKD1), which is detected at moderate levels in a subset of neurons in the anterior margin and in the sub-epidermal nervous plexus (Figure 3). We detected CAVII-1 expression first, then incubated in either 2% hydrogen peroxide (Figure 6A), 4% formaldehyde (Figure 6B), 100 mM glycine pH 2.0 (Figure 6C), or 100 mM sodium azide (Figure 6D) for 45 minutes to quench peroxidase activity, before finally detecting PKD1 expression. Peroxidase quenching using hydrogen peroxide has been widely used in planarians [21]. However, we find that while hydrogen peroxide does effectively quench peroxidase activity it also led to an increase in background fluorescence for PKD1 (Figure 6A). More recently formaldehyde fixation has been used in planarian multicolor FISH experiments [34], but while signal sensitivity for PKD1 was unaffected, there was clearly residual peroxidase activity from detection of CAVII-1 (Figure 6B arrows). Inactivation of peroxidase activity using low pH has been described in other systems [35, 36], and was effective at eliminating residual peroxidase activity from detection of CAVII-1 (Figure 6C). However, it also greatly reduced signal intensity for PKD1. We found that incubation in 100 mM azide was effective at inactivating peroxidase activity without reducing signal sensitivity (Figure 6D), particularly when determining coexpression of low abundance transcripts (Figure 7). While the utility of azide in quenching peroxidase activity has been examined [36], it has not been widely used for multicolor FISH or immunohistochemistry using TSA. One possibility for this is that it has typically been used at lower concentrations where it may be less effective compared to other methods. Our results suggest that use of high concentrations of azide to inactivate peroxidase activity could prove useful for multicolor FISH experiments as well as for immunostaining experiments using TSA in other model organisms.

Azide effectively quenches residual peroxidase activity in multicolor FISH. (A-D) The efficacy of several peroxidase quenching compounds was compared by performing multicolor FISH for Smed-CAVII-1 (expressed at high levels in protonephridia) and Smed-PKD1 (expressed at moderate levels in a subset of neurons). CAVII-1 expression was detected, then residual peroxidase activity was quenched by incubating samples for 45 min with either 2% hydrogen peroxide (A), 4% formaldehyde (B), glycine buffer pH 2.0 (C), or 100 mM azide (D). Finally, a second TSA reaction to detect expression of PKD1 was performed. Arrows indicate signal from residual peroxidase activity of CAVII-1 in the green channel. Scale bar: 100 μm.

Modified FISH protocol for determining expression patterns of problematic genes. (A) Maximum intensity projection showing enriched expression of Smed-hb (magenta) in ciliated protonephridia labeled by anti-acetylated α-Tubulin (blue) and Smed-POU2/3 (green). A subset of POU2/3 positive putative protonephridial progenitor cells in close association with ciliated protonephridia coexpress hb (arrow). (B) Single confocal section showing coexpression of Smed-CHD4 (magenta) and smedwi-1 (green) in intact planarians stained with DAPI (blue) and the chromatoid body marker, Y12 (yellow). Most chromatoid bodies lack CHD4 mRNA (closed arrows). However, there are occasional chromatoid bodies in close association with CHD4 puncta (open arrowhead). (C) Maximum intensity projection of a cell expressing Smed-nog1 (magenta) and Smed-nog2 (green). Scale bar: 10 μm.

Application of the in situhybridization modifications to detect difficult transcripts

With the improved signal sensitivity we were able to achieve with our modifications to the planarian FISH protocol, we sought to resolve the expression patterns of a few genes with unclear expression. The transcription factor Smed-hb (hb) is required for normal maintenance and regeneration of protonephridia [15, 38]. However, based on the published expression pattern it is unclear whether hb is expressed in protonephridia. Therefore, whether it acts autonomously in protonephridia or non-autonomously remains unresolved. We used hb as a representative problematic transcript for optimization of our FISH protocol, and found dramatically improved signal sensitivity particularly following formamide bleaching (Figure 1O). While we did observe broad expression of hb following chromogenic detection, we noticed staining of tubular structures consistent with protonephridial expression as well as stronger, punctate expression throughout the animal. To determine whether the tubular expression coincides with protonephridia, we performed FISH for hb and immunostained with anti-acetylated α-Tubulin antibody, which labels cilia in the lumen of protonephridial tubules [39–41]. We observed clear signal for hb surrounding ciliated protonephridia, consistent with hb function being required autonomously in protonephridial cells (Figure 7A). Scimone et al. (2011) have described a population of protonephridial progenitor cells defined by overlapping expression of protonephridial transcription factors [15]. To examine whether the punctate staining we observed for hb might represent protonephridial progenitor cells, we performed double FISH experiments with hb and several protonephridial transcription factors, including Smed-POU2/3, and found that a few of the hb-positive cells also expressed POU2/3 (Figure 7A). While we did observe expression of hb outside of the protonephridia, these results support the possibility that hb function may be required in protonephridial progenitor cells and mature cell types.

The planarian gene Smed-CHD4 (CHD4), is a homolog of the chromatin remodeling gene CHD4/Mi-2, and is required for the normal differentiation of neoblasts [22]. CHD4 has been reported to be broadly expressed and enriched in the central nervous system [22]. Consistent with CHD4 expression in neoblasts, mesenchymal expression is reduced following lethal irradiation, and CHD4 in situ hybridization of sorted cells results in labeling of a significant fraction of neoblast subtypes [22]. While these data provide compelling evidence that neoblasts express CHD4, we wanted to see if we could verify coexpression of CHD4 with other neoblast markers in intact animals. For this we performed multicolor FISH for CHD4 and smedwi-1. While we noticed broad expression of CHD4 throughout the animal, there was clear punctate expression of CHD4 in the cytoplasm of neoblasts (Figure 7B). The punctate, cytoplasmic localization of CHD4 RNA in neoblasts is reminiscent of transcripts present in ribonucleoprotein complexes called chromatoid bodies that are believed to be important sites of post-transcriptional regulation [42]. To examine the possibility that CHD4 mRNA localizes to chromatoid bodies we immunostained with the monoclonal antibody Y12 [43], which recognizes symmetrical dimethylarginine in proteins associated with chromatoid bodies [44]. While we occasionally observed CHD4 puncta (magenta) near chromatoid bodies (yellow) (Figure 7B open arrowheads), we rarely observed overlap between Y12 immunostaining (Figure 7B arrows) and CHD4 FISH signals. This observation suggests that the punctate signals observed represent subcellular localization of transcripts to cytoplasmic regions other than chromatoid bodies.

The expression patterns for several members of the noggin gene family in planarians have been described [23]. Smed-nog2 (nog2) is one of a few noggin gene family members whose mRNA distribution remained elusive despite confirmation of expression by Reverse Transcription-quantitative PCR. While knockdown of either nog1 or nog2 alone yields no phenotype, nog1 nog2 double knockdown leads to a dorsalization phenotype [45], further bolstering the likelihood that nog2 is expressed. In single FISH experiments we were able to detect nog2 expression (Additional file 2), and observed a pattern similar to that of nog1. We were curious to determine whether nog1 and nog2 are coexpressed, or whether different cells contribute either nog1or nog2 to regulate dorsoventral polarity. To examine this we performed double FISH for nog1 and nog2. Signal for nog2 was clear but significantly weaker than for nog1. We found only a small percentage of nog1-positive cells that also expressed nog2 in the body margin (Figure 7C). The more limited expression pattern for nog2 compared to nog1 could be real, or may indicate that expression of nog2 in some cells is below the limit of detection. Despite the latter possibility, our ability to at least partially detect gene expression for a gene that has been refractory to analysis highlights the utility of the modifications we have established for this protocol.


Mouse oocyte/embryo collection

All animal procedures were reviewed and approved by the Institutional Animal Care and Use Committee at the University of Nebraska-Lincoln and all methods were performed in accordance with relevant guidelines and regulations. For this specific study, CD-1 outbred mice had ad libitum access to water and normal rodent chow (Harlan Teklad, T.2918.15) they were maintained on a 12:12 dark: light cycle. At 6–8 weeks of age female mice were stimulated with 5 IU equine chorionic gonadotropin (eCG) and 5IU human CG (hCG) as described 39 . Cumulus-oocyte-complexes (COCs) containing germinal vesicle (GV) stage oocytes were collected 44 hours after eCG by puncturing antral follicles on the ovarian surface with a 27-gauge needle, while unfertilized MII oocytes were collected from the oviduct ampulla 16 hours post-hCG. To collect 1- and 2-cell embryos, eCG/hCG-stimulated females were placed with intact males of proven fertility overnight. Presumptive one-cell embryos were collected from the oviduct ampulla 16 hours after hCG stimulation. Two-cell embryos were flushed from the oviduct 1.5 days after mating.

Oocyte/embryo fixation and single molecule RNA florescent in situ hybridization assay (RNA-FISH)

Freshly isolated cumulus-oocyte complexes, MII oocytes and pre-implantation embryos collected from at least 3 CD-1 mice per developmental stage were fixed in 100 μL drop of 4% paraformaldehyde with 0.1% polyvinylpyrrolidone (PVP) for 20 min, washed through 3 drops of wash buffer (0.1% Triton X-100 and 0.1% PVP in 1 × PBS) and permeablized (1% Triton X-100 in 1 × PBS) for 30 min. After permeablization oocytes and embryos were placed in wash buffer for 10 min prior to hybridization steps. Each hybridization step was performed in solutions within one well of an Agtech 6-well Solution dish (D18, Agtech, Manhattan, KS).

Affymetrix ViewRNA Cell Assay Hybridizations

Oocytes and embryos were subjected to limited protease digestion using 80 μl of QS diluted 8000-fold in 1 × PBS. Following the 5 min protease treatment oocytes and embryos were transferred to 100 μL wash buffer for 10 min. Permeablized and protease-treated oocytes and embryos were subsequently transferred to 80 μL of probe-containing solution (proprietary ViewRNA ISH probe sets for murine Gdf9, Pou5f1, or Nanog (see Table 1) diluted 1:100 in QF diluent) and incubated for 3 hours at 40 °C. For the studies presented, oocytes and embryos were co-hybridized with probes for Pou5f1 and Nanog while single hybridizations were performed using Gdf9. Negative control cells were hybridized in QF diluent with no probe set added. Due to the specific gravity of the probe containing solution, oocytes and embryos tend to float and become transparent. It was crucial that oocytes and embryos were fully submerged during the entire 3-hour incubation. Following probe hybridization, oocytes and embryos were washed and then subjected to sequential hybridization with pre-amplifier DNA, amplifier DNA and fluorophore (Gdf9, LP1-550 Pou5f1, LP6-650 and Nanog, LP4-488). Each of the proprietary hybridization DNAs/label were diluted 25-fold in the provided diluent. Hybridizations with pre-amplifier, amplifier, and fluorophore were performed at 40 °C for 30 min each.

Advanced Cell Diagnostics RNAscope Hybridizations

Oocytes and embryos were subjected to limited protease digestion using 1 × Pretreat 4. After 5 minutes, oocytes and embryos were transferred to wash buffer for 10 min. Permeablized and protease-treated oocytes and embryos were subsequently transferred to 80 μL of probe-containing solution (proprietary RNAscope 3-plex Positive Control Probe Mm containing M. musculus Ubc, Ppib, and Polr2ra (see Table 1) combined as described by the manufacturer) and incubated for 2 hours at 40 °C. Alternatively, cells were hybridized with Negative Control probe containing B. subtilis DapB (see Table 1). Due to the specific gravity of the probe containing solution, oocytes and embryos tend to float and become transparent. It was crucial that oocytes and embryos were fully submerged during the entire 2-hour incubation. Following probe hybridization, oocytes and embryos were washed and then subjected to sequential hybridization with pre-amplifier and amplifier DNA (Amp1-FL, Amp 2-FL, and Amp 3-FL) and fluorophore (Amp4A ltB Ubc-647 nm Ppib-488 nm Polr2a-550 nm DapB -550 nm, 488 nm, and 647 nm). Hybridizations with pre-amplifier, amplifier, and fluorophore were performed at 40 °C for 15–30 min each as indicated by the manufacturer.

Following each hybridization step (regardless of the kit), oocytes or embryos were transferred through 3 wells of wash buffer. Oocytes and embryos were subsequently counterstained with 4′, 6-diamidino-2-phenylindole (DAPI) for 20 minutes prior to mounting on 25 × 75 mm microscope slides (Gold Seal®, 3039) in 12 uL ProLong® Gold Antifade Mountant reagent (P36934, ThermoFisher Scientific) and 25 × 25 mm coverslips (48368 084, VWR, Radnor, PA).

Confocal Imaging and mRNA quantification in individual oocytes and embryos

Hybridized oocytes and embryos were imaged using a Nikon A1 “LSCM” on a Nikon-90 laser scanning confocal microscope at the University of Nebraska-Lincoln Center of Biotechnology Microscopy Core with image capturing assistance provided by Dr. Christian Elowsky. Sequential imaging was performed to avoid nonspecific fluorescence detection. Appropriate filter sets were applied based on the fluorophore used for each transcript (Gdf9, Polr2a, and DapB 550 nm Pou5f1, Ubc, and DapB 647 nm Nanog, Ppib, and DapB 488 nm) and Z-series sectioning performed. Individual Z-section images were visualized with NIS-Elements 4.4 image program and exported for compatibility with Image J. To count individual fluorescence signal indicative of a single mRNA molecule, images for each section were maximum-projected and stitched together in Image J Fiji using the Grid/Collection plug-in 27 to form a composite image (1.45 S, Wayne Rasband, NIH, USA). Individual spots of fluorescence were located and counted in each composite image using the software program Localize which was written in Interactive Data Language (ITT Visual Information Solutions) 40 . Specifically, the Localize program was run using each composite image and the default threshold of 10.0 photons and band pass threshold of 400 (Fig. 3A) the output of the program was the number of spots counted.

Traditional RNA isolation, reverse transcription and droplet digital polymerase chain reaction (ddPCR) analysis of target transcripts

RNA was isolated from a pool of 15–20 oocytes or embryos collected from at least 2 CD-1 mice per developmental stage using Tri Reagent (Sigma-Aldrich) according to the manufacturer’s instructions. Isolated RNA was reverse-transcribed using MMLV-RT as previously described 41 . Complementary DNA (cDNA) from oocytes and embryos were diluted 2-fold prior to combination with 1XQX200 ddPCR Evagreen Supermix (BioRad Laboratories), which includes a proprietary SYBR green fluorescent dye and RNA polymerase, and 100 μM of gene specific primers (Table 1). In addition, primers were combined with either no template (PCR negative control) or synthetic-produced gBlocks Gene Fragments (10 pg/μL, Integrated DNA Technology, Table 1) which represented the PCR positive control. Each sample was emulsified by the QX200 Droplet Generator (BioRad) resulting in 20,000 droplets containing the Evagreen supermix (Bio-Rad), cDNA template with or without the targeted sequence, and target cDNA primers per reaction tube. Forty rounds of PCR amplification were performed using the C1000 Touch Thermal Cycler (Bio-Rad) and the number of droplets positive and negative for fluorescence for each sample was measured using the QX200 Droplet Reader (Bio-Rad). The copy number of target transcript per μl of cDNA was quantified using Quantisoft (Bio-Rad) and the number of target transcripts μ per oocyte or embryo calculated as follows:

Statistical Analyses

Comparison of Gdf9, Pou5f1, and Nanog mRNA abundance between in vivo produced GV-stage cumulus oocyte complexes, MII oocytes, 1-cell embryos, and 2-cell embryos was performed using GraphPad Prism 7.0 (GraphPad Software, La Jolla, CA). For each comparison, one-way ANOVA was performed followed by Tukey’s multiple comparison post-test. Data were presented as mean ± SEM. Differences in mRNA abundance between each oocyte or embryo stage were considered significant at P < 0.05.


Introduction: Developed by Advanced Cell Diagnostics, RNAscope ® in situ hybridization technology enables detection of a target RNA in a cell-specific manner on formalin-fixed paraffin-embedded tissue sections and represents a good alternative to immunohistochemistry. The goal of this work is to illustrate an optimized protocol of the RNAscope technology to detect target genes in various human organotypic culture models (nasal, small airway, and gingival). These culture models retain the three-dimensional structure of native epithelium, mimic in vivo morphology and human physiology, and can be used as alternative sources to animal testing.

Materials and Methods: After fixation and processing of five replicates of the three different organotypic cell cultures, the tissue morphology was checked by hematoxylin and eosin staining. The RNAscope protocols were optimized based on three crucial parameters: heat pretreatment, enzymatic digestion, and signal amplification. Digital images of the RNAscope stained slides were generated using the Hamamatsu NanoZoomer 2.0 slide scanner, and images were quantified using a custom-made plugin on Definiens Tissue Studio software (Definiens AG, Munich, Germany).

Results: The tissue morphology demonstrates optimum fixation and processing for samples, while the optimized protocol for RNAscope shows preserved RNA with staining on the positive control probe with score ≥2 and no staining on the negative control probe with score <1.

Discussion and Conclusion: RNAscope combined with organotypic cell cultures is a promising tool to better understand cell-specific RNA expression while implementing 3R (replace, reduce, and refine animal testing) principles.

About the authors

Steven Westra

Steven Westra is a renowned antibody staining consultant with over ten years of experience in the immunohistochemistry industry.

Geoffrey Rolls , BAppSc, FAIMS

Geoffrey Rolls is a Histology Consultant with decades of experience in the field. He is a former Senior Lecturer in histopathology in the Department of Laboratory Medicine, RMIT University in Melbourne, Australia.

James Anderson , Global Marketing Manager

James Anderson is a Global Marketing Manager at Leica Biosystems with experience with histology and scientific, technical, and marketing communications.


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