Staining - Science method

We are isolating hepatocytes from mouse liver and trying to culture them in 6 or 24 well plates. The isolation is carried out according to the protocol and plating is done using William E medium with 10% FBS and 1% PEST. We use to coat the plates with collagen 1mg/ml. But after 3 hours and more the cells are loosely attached to the plates. We are sure these cells are hepatocytes because we stained them with the appropriate markers. Maybe something is missing in the medium which is essential for attachment?

I have prepared an oil in water nano emulsion and would like to visualise it for confocal microscopy, however i dont have any fluorescent compounds in my emulsion. I know that nile red is a lipophillic dye and would like to know how to stain (protocol with concentrations) it so that i can visualise the droplet morphology better using confocal/fluorescence microscopy?

I work at the departement of pathology, EM-lab and we are on the look out for a replacement/update for the old Pyramitome 11800 we still use.

The one we have now is quite old, but the working angle is perfect for searching for glomeruli in Epon-embedded kidney-tissue. We have not found a replacement, and are wondering if anyone knows of one.

We have the Leica EM UC7 ultramicrotome, but the working angle is not ideal for the purpose of searching for glomeruli. We would have to "randomly" trim and stain to see if we have glom, which would take too much time and is too uncertain.

The added picture is the type we have.

Would appreciate any advice and tips.

Thanks

I am working with blood and A549 cancer cell. My experiment design is where I will stain the cancer cells with CellTracker CMTPX, and also stain the blood (lysed blood sample with a few residual RBC) with hoechst separately and then spike the stained blood sample with the stained cancer cells.

There are no issues with the dye used for cancer cell but the hoechst always stains the cancer cells a dull blue as well after I spike the blood with cancer cells. I have tried to wash the stained blood cells 5 times in PBS to prevent this stain contamination but it keeps happening. Is there any way to prevent this?

Hi,

I am looking for a viability marker to stain alive white blood cells and still working after fixation.

Here is my workflow for you to better understand (and tell me if my asumptions are wrong).

1) I am studying white blood cells from total blood (EDTA tube).

2) I take a certain volume of blood and had a buffer with fixative agent during 8 min (blood is diluted in 1:1).

3) Then, I make the blood go through a filter to stick fixed cells on it.

4) Finally, I stain the filter with May-Grünwald Giemsa - MGG (resulting in colors from pink to blue with purple) to see the nucleus, cytoplasm et identify my cells under microscope (brightfield).

My idea is to be sure that my cells I identify by MGG were well alive before the 8min fixation.

--> So, I would love to have a viability markers that I could identify in brightfield (so more a dye?), in different colors other than pink/blue/purple and that is not toxic for cells (no apoptosis or death process). If you have an idea.. let me know please :)!

I have made some research and here I found :

Thank you for you returns! :)

I am manually counting neurons in specific brain regions with bright-field stainings and I was wondering if anyone has a way to automate this? I am using the software Aperio ImageScope now, but am open to other suggestions! Thanks in advance!

I'm getting a problem with my hydrogels. As I wash my sections more rigorously, more hydrogel will wash away. However, the more you wash the slides, the more ECM and cells wash away as well.  

Thank you

Dyes for live-dead bacteria staining

Hi guys, I have some ceramide and dihydroceramide (d18:0/14:0) dissolved in methanol : chloroform=1 : 1. I tried to run TLC for both of them and stain the plate with Iodine and potassium permanganate. The band for ceramide is very clear but the band for dihydroceramide is always invisible. For the amount I used 10 micromoles for both of them.

Hi, I'm a master's student in biotechnology and I'm having trouble visualizing plasmid bands during extraction. I'm working with the plasmid pET-32a in *E. coli* DH5α. I've tried both kit-based and manual extraction methods, but I can't see any plasmid bands. I've run the electrophoresis at 90 volts for 40 to 45 minutes (1% agarose gel) using Safe stain for staining. After using the kit extraction method, the Nanodrop showed a concentration of 162 ng/µL. The DNA ladder shows clear and distinct bands, but the plasmid does not. Does anyone have suggestions on what might be going wrong or how to improve the visualization of plasmid bands?

Hello, I am trying to study synapses and try to stain synaptophysin and synapsin on postmoterm rat brain tissue. But so far have completely no siangls under confocal (I am expecting to see punctas).

Rat brain tissue is kept in 4% paraformaldehyde for over 3 years.

My protocols invoves 10% serum for preblock and 1% serum for antibody incubation. PH7.4 PBS buffer washes between steps and 1 night first andtibody and 1 night second. Dilution factor range from 1:200-500 for primary and 1:200 for secondary. I tried antibodies from Milipore and Fisher as how other paper used them.

I also tried staining PV for positive control and I see perfect singals (so the tissue should be fine).

Do you have any suggestions where the problem should be? or any suggestions of change s in antibodies use or protocol changes?

I just started staining slides for IF and have been seeing some cloudiness form when I prepare slides using thawed WBCs from Bone marrow samples. When I use fresh (not previously frozen) samples, I don't see this weird fogginess. I was wondering if anyone else has run into this issue before?

I've included pictures of a fresh stain versus a thawed stain here.

Can I substitute SSC buffer for PBS in propidium iodide procedure for staining of cells? Also, can I PI stain cells without fixing them, or is that not an option?

Hello everyone,

I have a question concerning the CellTracker Red fluorescence dye. Lately, our cells are dying when stained with CTR and then live imaged (we do timelapses over several hours, imaging ever 10-30 minutes). All the controls (stained, but not imaged and vice versa) seem fine, so it is (so far) just the combination of staining and prolonged imaging which kills the cells. As a final working concentration, the cells recieve 11 µM CTR for 45 min at 37 °C.

We are thinking of testing another dye to see if the fault lies with the imaging itself but that has not arrived yet.

Does anyone have any experience working with CTR and come across similar problems? Any suggestions would be welcome!

I have red blood cell coated with recombinant antibody. This antibody has 6X his-tag. we want to perform immunocytochemistry on RBCs. Which stain can we use?

I am trying to evaluate the cytotoxicity of my effector cells against an adherent target cell line. I plated 10,000 target cells and incubated them overnight to allow them to attach. I then stained them with 15uM calcein AM in complete medium and incubated the cells for 45 min at 37C. I washed three times with complete medium, then added my effector cells at a 1:1 ratio. I had a negative control (target cells alone) and a positive control (target cells with lysis buffer). I had two separate plates - after 4 and 24 hours, I transferred the supernatant to a black opaque 96 well plate, and added lysis buffer to lyse any effector cells that might've uptaken calcein. I have calcein AM blue, so read at 360/449 nm. When I read the signal, I saw very high background signal in my negative control wells, and did not see much difference between my experimental wells and positive control. Could this be due to insufficient washes? Or is there something else contributing to such high noise?

I want to stain my bacterial cells to highlight the nucleus on the surface where my protein is attached. I have tried using Hoechst, but the signal in flow cytometry was too weak, resulting in a low cell gate. Is there a better dye or an alternative technique to achieve this visualization?

I am trying to determine apoptosis in mouse liver tissues using the Elabscience TUNEL In Situ Apoptosis Kit (HRP-DAB Method) (Catalog no. E-CK-A331). In my first experiment, I was able to identify apoptosis in the tissues and get the ideal staining. Even though I did not change anything in the protocol, the next day, on my second try, I was unable to get staining in the tissues. What do you think could be the reason for this?

I’m losing my cell-embedded hydrogel discs during the washes needed for staining, which is causing me to lose my entire cell samples for imaging assays.

Any suggestions would be greatly appreciated please!

Hi every one, this is my first time doing luxol fast blue. My sections are mouse spinal cord-PFA 4% fixated-frozen-cryo-sectioned at 20 um. after 16 hours of LFB in 60 degree, I did not see any strong staining. After first differentiation, I look at them under microscope and they were weak and it was reverse( gray matter was blue and white matter was more clear)- I differentiated them second time and it was all gone.

What are your thoughts?

I am trying to stain spinal cord after cryo-sectioning.

I fabricated microbeads using hydrogel (20-50 micrometres) to mimic human cells' size and mechanical properties; I want to stain them to be able to recognize them under an optical microscope. Which staining should I use? I don't have confocal microscopy at the moment, so I think fluorescence inks will not be helpful here; I am using the microscopic camera to view those beads

Staining - Microbiology

I am recently observing DAB staining fail-regions (red circles or red arrows on pictures) on my DAB stained slides. Does anyone have ideas why I am getting this?

I can see only hematoxylin stained cells even in DAB positive area. But sometimes both DAB and hexatoxylin stainings were not stained in the fail-regions.

I used Leica BOND RX automated machine plus DAB staining kit for the DAB staining.

I baked the slides at 65 °C for 3 hours, so I don't think the fail-regions are because of not melted paraffin.

Moreover, the fail-regions are not seemed bubbles, because the shape is different from that of typical round bubbles(blue arrows in DAB-fail 3).

Does anyone experienced like this result? or has seen or solved the issue?

We have been trying to optimize IHC. We are using citrate buffer ph:6 for antigen retrieval in microwave oven for 20-30 minutes. Giving 1 hour incubation for primary antibody and 40 minutes for secondary antibody. But we are unable to get results. Hematoxylin staining is working fine but DAB is not going inside cells giving these brown spots. Attached are the images of recent try.

Thanks in advance.

I am encountering a significant issue in my experiments with U87 cell spheroids, using DiOC18 (D275) staining for fluorescence. Initially, the results seemed promising, as the expected green fluorescence was observed. However, upon analyzing the samples using the red filter, I noticed red spots within the spheroids, which likely indicate DiOC18 aggregation. This makes it impossible to proceed with propidium iodide (PI) staining, as these aggregates could result in false positives. The DiOC18 was dissolved in DMSO and left under agitation overnight. The stock solution was prepared at 2 mg/mL, and the working solution was adjusted to 20 µg/mL.

I want to treat fibroblasts with 24h MG132 and then look a the amount of cell death. However, I want to correct for the amount of cells in total, because they also prolfirate in 24hour.

My supervisor suggested that after 24hour I stain my cells with HOECHTS and PI and then quantify with imageJ. The amount of dead cells can be calculated with:

x amount of HOECHTS positive cells/ x amount of PI positive cells.

However, HOECHTS also stains apoptotic cells. I do not know if this will interfere with my results. Do you think that after 24 hours most apoptotic cells are aleady dead anyway (or at least a part, normalizing the results)?

Or is this an unreliable method?

I'd like to use the confocal to track spermatogenesis in various coral colonies  over the season without having to do too much histology.  But, I'm not sure if this is possible with the confocal/fluorescent dyes.  Side bonus... corals and their symbionts auto-fluoresce green/red respectively.  So, looking for something in the blue (400-450 nm) and yellow (565-600 nm).  If it's possible for one to target spermatogenesis and the other to track oogenesis, then that's just lovely.

Good morning,

I am working on rats and currently testing some coordinates for NMDA lesion. Usually for NMDA lesion we wait approx a week and we perfuse with PFA 4% before vibratome cutting and a simple Thionine staining (or NeuN but here I do not need that). Because we want to win some time and also because we want to use the cryostat, I was wondering if it will be possible to visualize properly the NMDA lesion on fresh tissue and also if with the damage it's not too risky to not perfuse the tissue before freezing and cutting it. I've already done coordinate pilots with indian ink and Neutral red staining on fresh tissue cutted with cryostat but NMDA is the next step.

If anyone have already done that or has a reasoned opinion, I will really appreciate.

Thank you!

We culture cells on a microchip with a membrane that has autofluorescence and after few hours the stained cells start fading I am searching for a cell tracker dye with strong intensity to make it easy to track cells in this high noise background. 

I took some blood from a B6 mouse that was hit with tumor cells and did a Richter's Giemsa staining. This is the first time I did Richter's Giemsa staining, and I couldn't identify the cells in it for sure. Is there anyone who could help me with this or tell me the possible problems with my staining results

I want to visualize the bacterial colonies grown inside a closed system made of transparent polymer. For this, I am planning to use inverted fluorescent microscope. Please suggest what magnification will be best 40X or 100X? Also, kindly share the fluorescent dyes to be used to stain bacterial membranes?

My lab received some mice heads that have been (and still are) in PFA for more than 2 years. What do I have to do to be able to extract the brains? And is it possible to freeze this tissue and use it for immunochemistry stainings?

Hey. I have a couple of methyl green stained tissues but the staining was not very good and I'd like to de-stain and try h&e. I've seen a couple of H&E de-staining protocols and I'm not quite sure if they are useful in this case. Do you know any methyl green de-staining protocol?

Thanks

Dear all,

I'm putting together an antibody panel to characterise airway epithelium.

I've tried two different p63 antibodies which show mostly unexpected cytoplasmic staining. I would expect 15-30% of epithelial cells to be positive with nuclear staining.

Has anyone else come across cytoplasmic staining in airway epithelium?

Details as follows:

FFPE section of lung tissue. Antigen retrieval done on Leica Bond Rx, blocked in BSA with Dk serum, Triton X-100 and Tween 20.

C1 (blue): DAPI

C2 (green): Ms x p63 (ab735, 1:100)

C3 (red): Rb x p63 (ab124762,1:200)

C4: merge

Thanks!

Hello everyone, I am currently working on Nile Red staining and fluorescence microscopy to confirm the presence of PHB in microalgae. Both lipids and PHB in microalgae can be stained by Nile Red, how can I distinguish between them? Additionally, what methods can be used to remove Nile Red staining from lipids?

Hi guys. I was wondering if you guys have any suggestions on staining live cells for long time (ideally at least 5 hours)? I want to at least define where the cell boundary is, and hopefully it should be a red dye so it will have less crosstalk with my another green dye. I have tried several dyes but none of them satisfies me:

I am running out of ideas now. The only option left is expressing fluorescent proteins tagged to other membrane proteins. CellBrite steady looks like another great choice since it is expected to retain signal at the PM for more than 24 hours. It might work but just could take too much time. Would appreciate it if anyone have experience on this topic ;)

Until I create my strains with a GFP-tagged protein of interest, I've been doing immunofluorescence on yeast to look at this protein of interest examine its location.

After primary incubation, washes, secondary incubation, and more washes I subject the yeast to 10uL of DAPI at 1:1000 for 5 minutes at RT. Then, I wash 2 x 5 minutes with PBST (0.1% Triton-X 100).

Sometimes the DAPI staining produces a halo-like effect in addition to regular nuclear staining. There also seems to be a lot of cytoplasmic staining in some cells. I do not believe the nucleus was compromised. Does anyone know the cause of this?

when I use SYBR Green II nucleic acid gel stain (1:10000 in TBE buffer) with mIRNA, why fluorescence decrease with increasing the concentration of mIRNA? I tried many times but same. I select the excitation is 579 nm and emission is 500-600 nm.

Hello everyone,

I'm currently working on immunofluorescence staining experiments, and I've noticed a consistent issue. Regardless of the primary antibody I'm using, I always observe higher background staining in the epidermis compared to the dermis.

I'm careful with my blocking steps and have tried different blocking agents, but the problem persists. Has anyone else experienced similar issues, or does anyone have insights into why the epidermis might show higher background staining? Any suggestions on how to reduce this would be greatly appreciated.

Thank you in advance for your help!

When I run this protocol, after passing the stains, the sample looks fine, but the stain off during dehydration and mounting. I do 3 stations in absolute alcohol and place in xylene. I have been following this protocol for years, but recently this problem has occurred.

I would like to stain live bacteria with plasmids encoding fluorescent-labelled proteins. I bought, for instance, one plasmid encoding a blue fluorescent protein (https://www.addgene.org/14891/). I got the bacteria, but when I put them under a fluorescent microscope, I saw no signal at all.

A I missing something? Do these kinds of plasmids need to be activated? (I understood they were constitutive). Are there better plasmids for the job?

Thank you

I am staining cells for a nuclear gene regulatory protein (red in attached image) which should only exist in the nucleus. I expect to see the gene regulatory protein forming condensates within the nucleus. Instead, I am seeing a ring of stain around the nucleus. Is DAPI interfering with my stain for the nuclear protein? Cells were permeabilized. So why am I seeing this ring?

Hi, I performed western blotting for the 8 times. In two of them I used wet-transfer and the rest of them assayed with semi-dry transfer. I checked my buffers, systems, gels and everything. I used two different primary antibodies which one of them is surely working. In the first 5 assay, I observed signal from the antibody that we know it is working. after we took images, I stained the membrane and observed protein bands on it. but now I can not observe any signal neither of them. and there is any protein bands on membrane. what could go wrong?

I‘ve been using an unconjugated mouse anit rat anitbody to stain stem cells. The antibodies are scarce so unconjugated format is the onyl one we could obtain. we used a secondary antibody FITC. For controls I used an isotype fitc mouse Ig1.

I determined the positive cell populations from the isotype control.

However, was it necessary to add a secondary antibody(FITC) only control to determine the positive cell populations?

Hi! I was staining my tissue with both PI and Calcein AM dyes, but I realize a lot of cells were stained by both PI (red) and Calcein AM (green), for example, the cell at the center of the attached image. I thought PI stained for dying cells while Calcein AM stained for live cells.. so I am curious to know if those cells are actually dying or alive (or both...??).

Also, some of the red signal was really blurry.. could it because those cells at the late stage of dying already (DNA was fragmented and released into interstitial space?)?

Thank you in advance!!

Dear colleagues, help me determine what kind of artifact this could be? After fixation in 10% formalin, dehydration with ethanol, paraffinization and routine staining with hematoxylin and eosin (Mayer's hematoxylin). I find that in some sections of rat liver, black nucleus, dots (from 3 to 30 pieces) are found. This does not look like the images of formalin pigment, hemochromatosis or other pathologies that I managed to find. I will be grateful for any help!

I need assistance quantifying migrated PBMCs using a transwell assay. PBMCs (50 000 cells in 50 µl) were seeded in the upper chamber of a Corning HTS Transwell 96-well permeable support plate (insert pore size 8 µm) in serum-free RPMI 1640 medium with GlutaMAX. The lower chamber contained medium supplemented with or without the chemoattractant, 10% FBS, and the plate was incubated for 24 h. After incubation, the non-migrated cells were removed from the upper chamber. The migrated cells that adhered to the underside of the inserts were fixed with 4% paraformaldehyde for 30 min and stained with 0.2% Crystal Violet for 10 minutes, and rinsed with distilled water. The stained PBMCs were imaged to assess the number of cells that migrated through the inserts. Since PBMCs are semi-adherent, the non-adherent cells migrated to the bottom of the wells, where images were also taken for quantification. While I can manually count the Crystal Violet-stained adherent cells in ImageJ using the cell counter, counting the non-adherent cells will be more challenging due to their more concentrated distribution at the edges of the wells (see image), especially in the chemoattractant group where more migration is expected. Any assistance on how to quantify this in a quick and easy way would be greatly appreciated! Please note that some cells do adhere to the bottom of the wells, making it time-consuming to harvest each well of a 96-well plate and count the cell suspension using a Countess. I have also tried Crystal Violet staining in the bottom wells to measure the absorbance with a plate reader, but the results are inconsistent, as some cells are washed away during the process.

We do peripheral blood chromosome analysis, in the last month we could not get G band, at the same time we got bands on chromosomes that stained for C band. I do not know if the increase of humidity was the main reason. But I will write the protocol to recommend me change some details that may help to get the G-bands again.

Preparation

- 0.075 g of trypsin powder dissolved in 100 ml of PBS, then heated in a water bath 56-58C.

- 85ml of D.water + 5 ml KH2PO4 + 5ml Na2HPO4 + 5ml Geimsa stain. pH =6.9

Start Staining

- Slides inseted in trypsin Jar at 56-58 C. for 40 seconds.

- Washing slides quickly in cold PBS for less than 1 second.

- Slides inserted in Giemsa jar for 3 min and 30 Sec.

- Twice wash quickly in D.water.

In order to view/ find the laid thrip eggs is there any methods is available, if yes, suggest me the methods.

I performed H&E staining on epididymal white adipose tissue, but after the staining, the tissue disappeared. I also processed a liver sample as a standard, and the liver sample was fine after staining. What could be the problem with the WAT samples?

Hi there! I'm hoping to to look at co-localisation of cytokines and markers of different brain cell types (e.g., neurons, astrocytes, microglia) in juvenile rat brains (postnatal day 28) that have been embedded in paraffin. Ideally, I would like to be able to image co-localisation, and also to analyse the morphology of glial cells that show co-localisation with cytokines.

I was wondering if anyone has any experience with the effects of different tissue slice thicknesses on the effectiveness of immunolabelling and morphological analysis. Most of the studies that do double/triple staining with paraffin-embedded tissue seem to use 3-5 micrometers, but the studies that perform morphometry (and only stain for 1 marker) seem to use thicker slices, e.g., 20-40 micrometers. Are thicker slices of paraffin-embedded tissue harder to stain for multiple markers?

Any advice would be greatly appreciated!

Hi, I need to make some counting of blue SABGAL stained senescence cells. All cells will be fixed and stained T75 flask, then trypsinization, making some cell suspensions and counting with Cellaca PLX. Is it possible to do something like this? If yes, what kind of dye and assay should be used?

I am looking to stain live Saccharomyces cerevisiae cells using DAPI (Thermo cat # 62248) but have currently been unsuccessful. Does anyone have a protocol to stain these cells without having to fix them using either DAPI or Hoechst, I want to be able to visualize the nucleus in live cell? Also does it matter if you use DAPI or Hoechst, is one better than the other?

Hello!!

I am doing histopathological analysis of kidney sections with PAS stain. I want to quantify the staining intensity of the sections. can anyone explain what is the correct protocol for using the color deconvolution tool in ImageJ software?

Thanks in advance,

Sreyasi!

I recently performed counterstaining with Mayer’s hematoxylin on mouse left ventricle sections. However, I observed dark spots, particularly in the infarct zone. Before proceeding with DAB staining, which might be compromised by these spots, could you please advise on how to address this issue effectively? And any advice to reduce the background of Hematoxylin statining

I am staining some brain sections stored in cryoprotectant that express a Histone H2B- GFP fusion protein that were generated ~10 years ago. I know I need to enhance signal with an anti-GFP antibody, but do I need any specialised H2B-GFP specific primary antibody? Or will anti-GFP do the trick?

Sections are free-floating 30um stored in cryoprotectant, initially fixed in PFA.

I have primary breast cells embedded in the hydrogel. When stained with EpCAM without the hydrogel, the staining was correctly localized to the membrane as expected! However, after embedding in hydrogel and using the same protocol for immunofluorescent, we are now seeing nuclear staining. Have you encountered this issue? Any recommendations on how to resolve it?

Hi all, I was just wondering if anyone has experience with multiplexing a mouse monoclonal primary and a rat primary. I'm trying to multiplex by incubating them in the same well but was told by a colleague to research the literature and find out if anyone has multiplexed with the same mouse monoclonal ab that we're buying and a rat primary. Their concern was background staining due to the species being alike (their secondaries could bind to the wrong primaries). We're using a mouse-on-mouse blocking reagent from vector. Does anyone have any experience with this or any recommendations? Thank you!

We tried both Kryo and Paraffin Embedding before cutting, and in both we had issues with autofluorescence in our COL1 scaffolds. Is it possible to avoid counterstainings/autofluorescence within the process of preparation of stainings (especially IHC with COL2,ACAN,DAPI) , and do you use some software/certain fluorescence markers to avoid this? Thank you for your help!

Hi,

I'm in a lab for 2 and a half years where we work on mouse tumors.

We have a protocol running for 7 years of tumor dissociation, staining, and facsing. We are using the cytoflex lx 5 lasers.

In the last 4 months, we have an issue that around 90% of the time we cant get a cd8a signal on facs.

We have no idea why- nothing changed, we're working with the same reagents from the same companies, we even tried opening a new vial of everything in order to make sure. Nothing changed.

When we can't get a cd8 signal, we're not getting it on any fluorophore.

On the time that it does work, the same protocol is applied.

Any ideas? Anyone had this issue in the past and solved it?

Thanks all!

Hello everyone,

I am currently using washed human platelets to stain Annexin V as a procoagulant marker. Additionally, I am staining with PerCP-CD61 to identify platelet cells. So far, I have tried using heat treatment (boiling in water for 20 minutes) and Triton to disrupt the cell membrane. Does anyone have other suggestions for the preparation of Annexin V positive control?

Thank you!

I want to know if there is any low cost method to identify an exopolysaccharide produced by bacteria is cellulose or other polymer.

Hi guys If anyone is currently working on aging cells, you guys would like to give me some advice. I'm testing against biomarker (SA-beta-Gal), I encountered a false positive in the control group and the senescence induced group had a worse stain. I have raised many questions about it staining endogenous lysosomes, also changing pH, osmolarity (fixative time)... I have tried all kinds of methods and followed the protocol as well but my results are not feasible..

Thanks a lot.

The organism used in the biofilm inhibition assay is Staphylococcus aureus. While performing the experiment in microtitre plate, I used 200 microlitres of (1:100 dilution of culture broth with absorbance 0.5). The incubation time was 24 hours. I could not observe biofilm formation at all even in the growth control when stained with crystal violet. Help me with suggestions as to how to improve the conditions for biofilm formation? Does increasing incubation time or using specialized broth help in biofilm formation by this bacteria?

I am doing an IHC protocol on a testes FFPE tissue slide. For primary Ab I am using Anti-Geminin antibody [EPR14637] from Abcam. For secondary conjugate I am using HRP. For chromogen I am using DAB. I am not co-staining the slides because I just want to look at geminin expression. The geminin ab is new and works well, same with HRP. I am following the general IHC protocol but at the end of the dehydration step and after mounting the slides, I am not seeing any staining develop. It's so weird, because I can see the reaction happening and the tissue getting darker with stain after washing and dehydrating them, the staining just vanishes.

Can anyone please help me with this?