A number of things to consider here: Firstly, there are major differences in the sensitivity of the two techniques, the way the data is displayed, and what constitutes a signal. Generally speaking, flow is the more sensitive assay, with data displayed on a log scale. In contrast, imaging is somewhat binomial: You decide what's positive based on what your eye sees, you set the laser power/PMT voltage/exposure time accordingly and then acquire. Even when images are collected in 16-bit format, most of the extended dynamic range looks black, and the signal you do see is pretty much linear. This means that signals that look middle-of-the-range by flow can still be black by imaging. To get a better idea of what signal is equivalent, you need to do a direct comparison between the two approaches using known fluorescence standards (e.g. rainbow beads) using the same settings as for your experiments. This would give you an idea what the limit of detection is for the microscope and what the equivalent signals are for each assay.

That being said, I thing that, in this particular case, the discrepancy might be due to transfer of "bits" of cells during the sample processing for flow. When you process a sample (mashing/digesting/scraping/spinning etc.), you can get transfer of cell debris from one cell to the other cells in the sample (presumably mediated by binding of surface proteins). When you subsequently run the samples through the flow cytometer, any bright "bits" that have come from your positive cells and bound to your negative cells will give a low-intermediate signal**. Due to this process, the presence of positive cells in a sample can actually raise the signal of the negative events, something that is extremely difficult to control for. This phenomenon occurs more egregiously depending upon the cells in question. It doesn't occur much when you're working with T cells, since they're quite sturdy. But it happens all the time when you do flow on macrophages and dendritic cells, which seem to be more fragile and lyse into pieces more readily. Although I've not had much experience with virus work, I would expect that infected cells would be some of the most fragile cells to work with and hence some of the worst offenders for lysis during processing.

**Keep in mind that cell "bits" still have the same density as the original cell, so the still spin down with your cells, which means you can't actually get rid of them from you sample.

Other differences might be filter differences as already mentioned, which could result in differential detection of background autofluorescence. Or, less likely, the enzymes used to release the cells for flow (if you are using enzymes) might unmask new cross-reactive epitopes, or induce variable binding of target epitopes, thereby changing the staining pattern.

My suggestion is that you prep and stain the cells for flow, but then cytospin them and look at them my microscopy. It should prove informative.

It may depend on the emission and excitation bands that you use on the microscope (are you using a UV lamp?) and the excitation in the flow cytometer? If the same than there may be a concentration effect?

A number of things to consider here: Firstly, there are major differences in the sensitivity of the two techniques, the way the data is displayed, and what constitutes a signal. Generally speaking, flow is the more sensitive assay, with data displayed on a log scale. In contrast, imaging is somewhat binomial: You decide what's positive based on what your eye sees, you set the laser power/PMT voltage/exposure time accordingly and then acquire. Even when images are collected in 16-bit format, most of the extended dynamic range looks black, and the signal you do see is pretty much linear. This means that signals that look middle-of-the-range by flow can still be black by imaging. To get a better idea of what signal is equivalent, you need to do a direct comparison between the two approaches using known fluorescence standards (e.g. rainbow beads) using the same settings as for your experiments. This would give you an idea what the limit of detection is for the microscope and what the equivalent signals are for each assay.

That being said, I thing that, in this particular case, the discrepancy might be due to transfer of "bits" of cells during the sample processing for flow. When you process a sample (mashing/digesting/scraping/spinning etc.), you can get transfer of cell debris from one cell to the other cells in the sample (presumably mediated by binding of surface proteins). When you subsequently run the samples through the flow cytometer, any bright "bits" that have come from your positive cells and bound to your negative cells will give a low-intermediate signal**. Due to this process, the presence of positive cells in a sample can actually raise the signal of the negative events, something that is extremely difficult to control for. This phenomenon occurs more egregiously depending upon the cells in question. It doesn't occur much when you're working with T cells, since they're quite sturdy. But it happens all the time when you do flow on macrophages and dendritic cells, which seem to be more fragile and lyse into pieces more readily. Although I've not had much experience with virus work, I would expect that infected cells would be some of the most fragile cells to work with and hence some of the worst offenders for lysis during processing.

**Keep in mind that cell "bits" still have the same density as the original cell, so the still spin down with your cells, which means you can't actually get rid of them from you sample.

Other differences might be filter differences as already mentioned, which could result in differential detection of background autofluorescence. Or, less likely, the enzymes used to release the cells for flow (if you are using enzymes) might unmask new cross-reactive epitopes, or induce variable binding of target epitopes, thereby changing the staining pattern.

My suggestion is that you prep and stain the cells for flow, but then cytospin them and look at them my microscopy. It should prove informative.

As Ben said above, FACS is much more sensitive than IF, so if you have obviously positive/negative cells in IF, you should see them when you FACS them. I imagine that you have nuclear staining in your IF and therefore are sure that you stain cells and not debris.

Are you sure you are gating properly in your FSC/SSC graph? Have you checked that you do not have any cell on the axis (top and right) that you may gate out inadvertently? if so, then you just need to change the FSC/SSC setting to get your cells back into the usual window.

I've seen new students making that type of error when using settings for very different cells than theirs.

Hope this helps, good luck - let us know when you've found the problem.

C.

From my experience, flow cytometry is more sensitive. If not, I will check 2 more points.

1) Prepare secondary antibody only negative control (deletion of primary antibody), not only the mock infected NC.This procedure confirm the abnormal secondary Ab aggregation. 2) Check the fluorescence of the flow sample with your eyes. If IF positive signal deleted from the dissociation step for flow, the antigen or the antibody easily removed from the dissociation.

Normally, flow cytometry is a more sensitive and more quantitative than microscopy. Are you sure you are using the right secondary antibody for flow cytometry? Depending upon the flow cytometer you are using, some dyes used in microscopy cannot be used in flow. You need to talk to the technician who manages your flow cytometer and ensure that. If you are using the right antibody then, the next thing to do is a titration to see what conc. of primary and what conc. of secondary you need to get decent signal of flow.

Are you doing IF on live cells or sections? When doing IF on tissue sections, the antigens that your antibody binds to may also be intracellular, which would not be picked up on surface flowcytometry labelling.