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Review best practice for confocal microscopy: from sample preparation through to data analysis, common pitfalls and problems, optimizing instrument set up and the advanced applications for which confocal microscopy can be used.
This webinar is ideal for anyone who is interested in carrying out confocal microscopy or is currently using confocal microscopy, as well as anyone who would like to better understand the technique and how it can be applied to their experiments.
Dr Ann Wheeler is from the Core Imaging Facility at Blizard, Institute of Cell and Molecular Sciences, Queen Mary University, London.
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Good day, ladies and gentlemen, thank you for standing by, and welcome to today's Abcam webinar: Principles and Practice of Confocal Microscopy in Life Sciences. Your first speaker today is Lucy Purser, Senior Events Coordinator. I would now like to turn the conference over to Ms Purser.
LP: Thank you, Nathan. Hello, and welcome to Abcam's webinar on Confocal Microscopy. Today's principal speaker is Ann Wheeler, Core Imaging Facility Manager at the Blizard Institute of Cell and Molecular Sciences at Queen Mary's University, London. Ann carried out her PhD work at the Ludwig Institute of Cancer Research in London, and obtained her PhD in 2005 under the supervision of Professor Anne Ridley. Her postdoc work was carried out firstly at the Scripps Research Institute, La Jolla, California.
Upon returning to England, Ann joined Imperial College, London. It was at Imperial College, in collaboration with the Institute of Cancer, she carried out a siRNA screen to identify and characterize novel proteins involved in regulating actin cytoskeletal dynamics in cell-cell adhesion. Ann has been the Core Imaging Facility Manager at the Blizard Institute since 2009. She has been part of publications in several leading journals and has won awards for her microscopy.
Joining Ann today will be Miriam, Product Manager at Abcam. Miriam completed her Biology Degree at the University of Barcelona and a PhD from the Vrije University in Amsterdam, after which she joined the MRC Laboratory of Molecular Biology in Cambridge. Miriam joined Abcam in 2008.
Before we start, as Nathan said, just a quick reminder that questions for the Q&A section at the end of the webinar can be submitted at any time via the Q&A panel on the bottom right hand side of your screen. Also, when you log-off from the webinar you will be redirected to a webpage where a downloadable PDF of the presentation can be found. I will now hand over to Ann who will start this webinar.
AW: Thank you very much, Lucy. That was very kind of you to give me such a lovely introduction. Today we are here to talk about confocal microscopy, and to discuss how to use that technique to the best possible advantage in your research. Confocal microscopy as a technique is there to take high-resolution fluorescent images of your cells, your tissues and anything else that you may be happening to use as a biological specimen, and it does this by eliminating out-of-focus slides.
So what would you need to be doing confocal microscopy for? Well, probably most of you may have some idea already, because you've been asked to do this in your research. Confocal microscopy has a range of applications, and this is just a small summary, but there are many more ways to do a confocal experiment.
You can use confocal microscopy for localizing or co-localizing different proteins, and seeing how they interact together. Confocal microscopy can be used for imaging two, three or four fluorescent stains and seeing which parts of the epitopes and proteins are interacting. It can be used to generate 3D images of your sample and so you can actually look at it in real space and see how molecules are distributed in the whole volume, or a whole organism. It's possible to use confocal for semi-quantitative measurements, and so you can use it to quantify protein expression, or to do comparative studies between proteins when you've mutated them, or if you've perturbed them using siRNA or drugs, or other inhibitors. Finally, what most people tend to use confocal microscopy for is to acquire very high quality images for presentation. So it can be really used to generate that punchy image that will get your data published, or get people to come and look at your poster when you're presenting it.
So when would one need to use confocal microscopy? Well, there are limits to what a confocal can do, it can be used to visualize structures between 200 nm and around 1 cm in size, which can be fluorescently labelled. This means it has applications in standard cell biology, tissue biology and also single molecule imaging applications.
So next, what I'm going to do is just give you a little bit of a summary of some of the areas that I am going to be talking about today, and some of the things that I'm hoping that you will have an opportunity to learn about in this webinar. First of all, we'll be discussing how confocal microscopy works, what's inside the machine and how come it generates these high-resolution images with no out-of-focus light? I'm then going to be talking about how the confocal microscope generates multicolor images, and maybe some of the pitfalls that one can fall into if you aren't careful with the set up. I'm going to talk about collecting the Z stacks, and there's 3D images in confocal and the considerations you need to have before you set that up. Finally, I'm just going to go through some of the questions that I routinely hear as a core facility manager, which people have, generally speaking, when they are trying to do their confocal images. Although, obviously, we do have the Q&A session at the end, so please, please do feel free to ask some more questions there.
First of all, what's the difference between fluorescence and confocal microscopy? Why would you want to do it? I'm hoping that this slide here would illustrate this, so, as you can see here, we've got an example of the structure inside the cell. As you can see, with a wide fluorescence image, it just looks very washed out and very yellow and you can't see much detail. However, when you look on the confocal you start seeing a lot of structure. You would maybe say the staining here and here are equivalent, but actually using the confocal you remove the out-of-focus light you can start seeing, no, there are a lot of different structures in here. There's also STED space here that's black, and so your sample maybe isn't as stained in the way that you would think it would be.
This is an image of some cell-cell junction proteins, and here you may get the impression that they're quite widely distributed, but actually when you look in confocal microscopy you can see it's quite discrete. So confocal microscopy can actually be used to remove some of the artefacts.
Perhaps the most telling image for why one would need to use confocal microscopy versus fluorescent microscopy, is this image of a pollen grain. With the wide-field fluorescence microscopy, because of the out-of-focus slides and the membrane around the center of the pollen grain, above and below it, the entire thing looks red. Whereas, actually, with a confocal microscope you section through the middle of it and you can see that inside there is this structure which is stained with a different antibody. That is, in many respects, why many people choose confocal as their tool of choice when acquiring their images.
So how is it that confocal images are clearer, or why is it that the confocal microscope is so complicated? A lot of people in facilities will be expected to go through some fairly rigorous training before they're let loose on the confocal and the reason is, really, as follows. So your wide-field epifluorescent microscope may look like a complicated instrument, but actually inside it's fairly straightforward.
You have a high-pressure mercury arc lamp that emits light broadly across the spectrum, so that it emits light of a range of different wavelengths, ranging from in the near-UV about 350 nm, all the way through to 800 nm. This light is shone into your sample and there's out-of-focus light as well as in-focus light there, and then it's reflected by the dichroic mirror down through the objective onto your sample. The sample obviously is thick, it's not thin, it's not one atom thick, in general. If it's a cell it'll be a few microns thick, and so what happens is then the light is emitted from the sample and you'll have the focus light from your point of interest, but out-of-focus light around it as well. This goes back through the objective and to the dichroic mirror up to your camera. So the image that the camera contains the in-focus light from your sample, and also the out-of-focus light, and that can be the cause of the haze in the image that you saw in the previous slide.
So how does confocal microscopy work? Well, essentially, this was designed by a chap called Minsky in the 50s, and it was a real breakthrough. He established that if you used these things called pinholes, which are basically small metal pieces of equipment with an adjustable sized hole in the front, you can actually use it to cut out the out-of-focus light. So you, instead of having a mercury bulb you have to have a laser, and that is because there's a lot of in-focus and out-of-focus light, but there's only a small amount of in-focus light compared to the out-of-focus light. So you need to use a high-intensity light source when you're starting to image your samples. So this is generally done from lasers of a defined wavelength, the laser light is shone in, reflected down to your sample in the same way via the dichroic mirror onto your sample. Now, of course, even the laser light going in is in-focus and when you get to your sample you will have a volume excited, and so the light that is emitted by your sample will contain some in-focus and some out-of-focus light. This will be reflected through your objective, which may create a few more aberrations in your image, and then it comes up to the detector.
Now, before the detector, this is the bit where the confocal really does its business, is there's a pinhole and the distance of the pinhole from the detector, and the objective has been very carefully calculated. So it is in the conjugate plane from the sample which is a focal plane, so this pinhole is in the confocal plane, as in the opposite to the focal plane. Now, if you know about what wavelengths of light that's being emitted from your sample, you know what to expect. You can also know how big the wavelengths of the light in-focus is, and you know that distance of the wavelength, so then you can set the pinhole up so it is exactly that length, or that size. That means the pinhole will cut out the out-of-focus light emitted by your sample, and will only have the in-focus light present; and that's great, because you only have your in-focus light. But, again, similarly to the light source in the first place, this means that most of the light is actually removed by the pinhole, and there's only a small amount of in-focus light that reaches the detector. So the detector needs to be more sensitive than an average camera and, in general, what people use is a photomultiplier tube, and so this will sort of collect samples point-by-point as the light comes in and then it will feed it back to the computer where the image is built up.
So to recap, what does the pinhole do? Its primary aim is to remove the out-of-focus light. The pinhole is adjustable, and that's important because obviously the size of the pinhole will depend on the objective lens that you use, and the wavelength of the light. So if you're using short wavelength light, then the pinhole will need to be fairly small because the wavelength is short. If you're using something a longer wavelength like red dyes, then the pinhole will need to be larger because the wavelength is longer. To make life easy for us, so a very helpful optical physicist decided to measure the pinhole size in Airy units, rather than expecting us to do all the math for ourselves.
So on your confocal it will say that a pinhole is measured as one, and that means that just the in-focus light is present, and the confocal microscope in general will know which objective lens you're using and what wavelength of light you're using; you will tell it that when you set it up. So it will be able to calculate for you how big the pinhole needs to be set, and you've got some power to adjust that as you choose.
So just to recap that why it is that you need lasers, you need a more intense light source, because you've cut out all of your out-of-focus light. So there's only the in-focus light, which isn't a great deal, and then similarly that means that the detectors, there's only the in-focus light. So you need a much more sensitive detector, and usually this is photomultiplier tubes, but sometimes this can be EMCCD cameras as well.
How does the confocal actually collect your image? What does it actually do? It doesn't just sort of shine the light and then it goes into the detector, it's more complicated than that. People who have got some experience of confocal microscopy will know that collecting a confocal image can take a little bit of time, compared to taking a wide-field image. This is how this happens, so basically in confocal microscopy you'll select your sample and bring it into focus, and the machine will sort of place a virtual grid over your sample. Each cell in the grid is called a pixel. You can decide for yourself how many pixels there are. The confocal default to certain sizes that you can actually select that, so you could choose how big or small the pixels are for yourself. Then what happens next is a confocal will scan pixel-by-pixel your image, as illustrated in the bottom side here by the yellow light, which is the laser light coming across scanning pixel-by-pixel across your sample, and collecting the laser light that's in-focus from that given pixel, this then goes to the detector and builds up the image.
Now, as you can see, as I'm clicking through the webinar, this is not a very fast process and so that's why your confocal will take a little longer to generate light, than your normal sample, and this point scanning is one of the major causes of issues in some confocal experiments. This is because a laser light is very intensely shining on a very small point of your sample for a period of time, and that can cause some of the fluorescent dyes that you've used to label your sample to photobleach.
So one of the more important things you need to consider when you're doing confocal microscopy, is how you prepare your sample in the first place. It's important that you use lights which are known to be bright, and the technical term for this is quantum yield. The quantum yield is defined as how many photons you get out of your fluorescent dye, compared to the photons that you get in? So if you get lots of photons out of your dye, it'll have a high quantum yield and it's ideal for using confocal microscopy, so Alexa Fluor® dyes, for instance, are very good for this.
Basically, if you want to improve your confocal image quality, then there's a couple of things that you can do. Generally speaking, when you start imaging on a confocal it will default to a fast-scanning mode which you use to optimize the laser power and the sensitivity of the detector. However, the speed of the point scanning of the laser can be varied, and the slower that you scan your laser across, the less noise that you get because, obviously, you're leading your laser to dwell on an individual pixel for a little longer. However, obviously, if you leave your laser sitting on a point like this, as described by my laser pointer, if you leave it for too long you end up burning a hole in it. So there's a bit of trade-off there, you may need to slow your laser scanning down a little bit, but too much and you'll bleach your sample. It's also possible to make the laser scan an individual pixel in your image more than once, and then display the average of this. This averaging again reduces the noise and increases the signal, and so by slowing down the laser speed and also increasing the averaging of your pixels, will increase the signal and decrease the noise producing a really beautiful image like this illustrated here.
So how would one optimize the image acquisition? Well, the first thing you need to do is check whether or not your image is saturated or undersampled, and on the left hand side here I'm indicating an image which is saturated/undersampled. Generally speaking, your confocal software there will be some button that you'll need to press to show whether the pixels are saturated and undersampled. Each confocal is different, each manufacturer has got separate software, so you need to contact your confocal's manufacturer, or somebody who's more experienced in using that confocal to find out which button that is. Generally speaking, the saturated pixels show up as red and the undersampled ones show up as blue, and it's best practice to have no pixels of the saturated or undersampled in your image, because then you're truly showing all of your data. So even if the black background is black, well, if it's black by eye then if it's not undersampled and it comes up black, then you're truly showing what's there. You're not excluding that data, and similarly if you've got some structures that are quite intense, you need to try, if you can, to get them within the range of the detector. So you're showing the full dataset and you're not hiding anything, that's the best scientific practice.
Make sure your sample is not bleached by the confocal, which can very easily happen. The best way to do it is to set the laser power as low as you possibly can get away with, and this may still be reasonably high, depending on your sample, but you need to try and minimize the laser power so it doesn't bleach your sample. The detector should be set at the appropriate sensitivity, so the specimen is neither oversampled nor undersampled, and so it's fully within the linear range of what the detector can do. The background, in my opinion, should be in the lineage range of the detector, but it should be black.
So now we move on to the next part of this seminar, which is preparing your sample for multicolor confocal imaging. As you can see here, we've got some excellent images of neurons and different small cells in a piece of tissue, which has been labelled beautifully for multicolor imaging; and it's possible for anybody to acquire images of this quality.
So when you're setting up confocal scanning of multicolor images, first of all you need to understand properties of the dyes that you are using. So you need to understand what the excitation or absorption spectrum of your dye is, and what the emission spectrum is as well. From this information you should know then which lasers you will need to use to excite it, and which detectors will need to in place, or which filters will be in place to detect the dyes.
Now, in the average sort of confocals, which are available on the market right now, there will be pre-sets for common combinations. So people who are imaging something with DAPI, something blue, something green, something red there will be a pre-set combination for that. Although it is important for you to understand a little bit more spectrally about what you're using, and I'll discuss that later. Generally, when a confocal scans multicolor images, you can set it to show each channel separately and then a merged channel, and that's important because you will need to align the detectors for each of the channels that you're imaging, not just the merge. So we need to make sure that the green, the red and the blue are within the linear range of the detector on the confocal when you're acquiring your data for best practice.
What will happen is in a merged image if the signal is from separate channels, is in the same pixel - if you remember, the confocal divides up your image into a pixel. So if the sample - there's a signal from two different channels in the same pixel, it will show up as yellow as defined here.
So just to recap before we go forward, so I think it's best if I review the fluorescence theory, and so, basically, fluorescence theory works like this. Your laser light shines in and you have your dye which is sitting in a ground state. The excitation laser comes in, the dye absorbs some of the light, it jumps up into a high energy excitation state and the problem is then with the fluorescent dye that it gets stuck. So it can't just drop back down to the ground state, it's got a problem and it's got to find a way of losing the energy somehow. So what it does is it vibrates, it knocks itself into the surrounding molecules and it loses some of the energy and heat a bit, and then it comes down to a slightly lower energy state, and it falls back down to the ground state. There's a difference between the energy that it needed to be excited, and the energy that it will emit when it absorbs. So the photon that needs to excite it will be of high energy, it will lose some of its energy and then the photon that's emitted will be of lower energy. So that means when you're doing any sort of imaging, you will tend to have one color light that's shining in to excite your sample, and then when you look down the eye piece what you'll see will be a different color, and that's why.
So this describes the excitation spectrum and what you need to be looking at. Something that I commonly have as a question in my kind of facility is, well, what is all of this then? Basically, a dye will not be excited at only one wavelength, it will be excited across a range of wavelengths. So this is Alexa Fluor 488® and, as you can see, it's excited maximally around 488 nm. It's also excited at 450 slightly and it's excited slightly at 520 as well. However, if you're shining in a laser light at 488 you're going to get about 75% of the molecules being excited by that light. Similarly, with a dye such as Alexa Fluor® 568 you see that when you shine in a laser light of 568 nm, about 60% of that dye will be excited at that wavelength.
What is useful when you're comparing the excitation spectras between these two different wavelengths, is if you look at the 488 dye and then you look at the 568 white laser, none of the 488 dye is excited by the 568 white laser, and that's quite important. You need to have some sort of separation there, and there are online tools that can help you do this.
More of a problem is emission spectra and so this is the range of wavelengths over which your dye will emit when it falls down to that lower energy level, and drops back down to the ground state. So the 488 laser light from the dye, will emit between 500 and actually all the way up to 650 nm ever so slightly, but by 550 most of the emission has fallen off. Similarly, for 568 most of the emission is occurring at 600 nm, but there's a bit at 550 and then it's emitting still at 700 nm, just not particularly much.
So what does that mean? That means that there is an overlap in the emission spectra here between the 488 and the 568 dyes. That will mean that when you're setting your microscope up, if you're not careful you might get some of the signal of the 568 coming through into the 488 channel. Now, this may look a bit complicated in theory, but the next slide I will show you in practice.
This is what it looks like, so this is an Alexa Fluor® 488 dye and this is an Alexa Fluor® 568 dye. What's happened here is that they haven't been careful about how they set up the light that they collected, so some of the light from the 488 has actually come through into the 568. So what you see in this channel is it looks like that this protein is everywhere. However, actually, if you're more careful about how you set it up, so you separate out the 488 from the 568 specifically, you'd see that the 488 is indeed in the middle of the cells. But if you are collecting only the light images by Alexa Fluor® 568, you will see that this protein is only present on the cell-cell borders, and ultimately in the middle of the cell.
So problems with not managing the bleed-through can really cause misinterpretation of the data, and that's quite a serious thing. So it's very important that you understand how you set up the filters, or the detectors on your machine to specifically pick up the range of wavelengths that you're trying to measure, and no more.
So how can the confocal help with this spectral overlap? Well, one way it does it is by sequential scanning, and so this is done so each channel confocal is set to collect, it's done separately, so it will do the green channel first and then the red channel. This is usually femtoseconds apart, it's very, very quick, so you don't have to sit there for minutes. Then when you're doing the sequential scanning, you can choose to scan either the whole image or one line of the pixels of an image, so you can see either the whole thing or have both channels build up line-by-line. It is possible to image using simultaneous scanning, and that's where all the lasers on the confocal are scanning at once.
The issue with this is obviously you get the bleed-through, but the advantage is it's much quicker; it just depends on what you're trying to do. One compromise is using mixed scanning, and this means that you scan using two lasers and detect using two detectors when you know that there's no spectral overlap between the channels. So one common combination that people use for this is dyes Alexa Fluor® 488, so dyes that are excited by 488 nm, and then Alexa Fluor® 633 or Alexa Fluor® 647. These two dyes don't interact with one another at all, and so it makes it very easy to detect both of these simultaneously without any bleed-through occurring.
So this is just an example of what the difference between sequential and simultaneous scanning is, and just to repeat this. The way you scan your images can give very different results, and it's important to check how you're doing it before you start. So here this image is just the Alexa Fluor® 488 dye, this image is just Cy3 and if you sequentially scan you can see that the combination of the two does give you the cells separated. However, if you simultaneously scan then they overlap, and so in the merged image which is D, it may look like the signal is co-localized but this is really an artefact for the incorrect confocal set up. So if you're studying co-localization of your proteins, it's essential that you set up sequential scanning to present the interpretation of data.
Also, when you're preparing your sample for confocal, and particularly when you're doing co-localization, you need to ensure that each epitope is separately labelled and has very spectrally separate excitation and emission spectra. The confocal has filters for your dyes and so they can be specifically separated in the light path.
It's also important for you to include certain controls if you want to be really sure, and really rigorous about the way you're doing your experiment. So you need to include some kind of positive control for the fluorophore that you're imaging, if you aren't sure whether the antibody's going to work, so maybe an antibody that does work or a dye that works.
You would need to include some kind of negative control for staining; imaging people will often just have the secondary on its own to see if the dye that's being used in a secondary antibody promiscuously binds somewhere where it shouldn't.
Several biological samples are autofluorescent, especially brain tissue where you have a lot of NADH and those flavins are very fluorescent. So it's useful to include a sample with no staining, so you can actually set up your detector so they're not collecting any of this autofluorescent background.
It's also advisable to set up controls for spectral bleed through, so you have only one epitope labelled and then you'd collect using confocal in both channels. So, say, for instance, you were looking for an experiment where you were looking at co-localization between a green channel and the red channel, you would need to have a control where you'd only got Alexa Fluor® 488 which emits in the green, and then image that and see what comes up in the red. If something comes up in the red channel you know that there's a problem with the way the machine's set up, and so you need to alter the parameters on the red channel detection, so with the green alone you see nothing in the red channel. You need to do that experiment the other way around, so with the red alone and being sure that you see nothing in the green channel.
People will sometimes also include an isotype control to ensure that the staining is really specific if it's something that's really unexpected. I would strongly advise people to ensure that their samples are collected using sequential scanning.
So now I'm going to hand over to Miriam who is going to present you a short quiz.
MF: As Ann mentioned, it is quiz time, so let's see how good your fluorochrome knowledge is. The quiz will appear in the right bottom corner of your screen, and you have one minute to answer these three questions. So here are your first two questions, and here's your next one. So thank you very much for taking part, the answers will be revealed before the Q&A session. I'm giving back the microphone to Ann, who will take you through the second part of the webinar, I would like to encourage you to submit more microscopy-related questions.
AW: Thank you very much, Miriam that was great, very helpful. So now we need to talk a little bit more about collecting some of those really beautiful three-dimensional images on the confocal. As you can see from this example here of a developing zebrafish embryo, it's possible for you to sample whole organisms in quite a lot of detail, and you'll be able to rotate them around and actually really see how the structures look inside the sample.
However, there's a couple of considerations that you need to make before you set up your 3D imaging. So if we go back to that theory that we discussed at the beginning of the session, the confocal pinhole moves the out-of-focus light from the X and the Y planes, so just one slat plane. Well, actually, it does a little bit more than that, it removes these out-of-focus lights in the axial plane or the Z plane, I'm going to call it in this talk for ease. So you can see here this is an example of your in-focus light and there are X, Y and Z components to it. The pinhole removes the out-of-focus light from all of these, and this means that it's possible to use a confocal microscope providing stage moves to image the whole of the volume of an object in 3D without any out-of-focus light present.
One thing I will describe now, as it's commonly used terminology, is the acquired confocal X, Y and Z section is called an optical section. So the optical section is not completely flat like a thin piece of paper, it's actually more like a cube in a shape. Thickness of the optical section is determined by three things: numeric aperture of the objective, which is how much light is left through it; the confocal pinhole size, which is how much in-focus light is there; and the wavelength of the light. The numeric aperture of the objective is something that describes how well the objective gathers light with a specimen at a fixed distance. Every single objective will tell you what its numeric aperture is on it, so this is a generic objective here.
You can see the magnification is described there. This is a numeric aperture and so the higher that number the better at gathering light the objective is. You need to find out a bit about the numeric aperture of your objective to find out how good it's going to be at collecting optical sections before you start your 3D imaging experiments.
So to collect your confocal volume, you need to have ideally as much of the information as you possibly can about the image. So the optical sectioning must be ideal in the X, Y and Z axis, so you need to find out for your objective how thick your optical section is going to be. Then you need to set the increment that the stage moves to be pretty much the same as the optical section to collect all of the light that's there, so you can produce your 3D image, which is truly representative of what it is. That information you may not know off the top of your head, but your microscope manufacturer may be able to tell you.
As an example here, I've got some information about the sort of objective you have, this numeric aperture and then the optical slice thickness, given that the pinhole is one. So you can see for a 10x objective with a numeric aperture of 0.3, which is quite low, the optical slice section is actually quite fat, it's 11 µm. So, obviously, if you've got something very thin then you probably aren't even going to need to do a Z stack, and what will present in your one slice is all the information that you need about your sample. If you go up to higher power objectives, as high numeric apertures, you can see the size of the optical slice drops off. So when you're using a 63x oil objective, if you have a high numeric aperture such as 1.4, the optical section thickness is 0.3 µm. So then, of course, that depends on how big your sample is and if it's large like that zebrafish, you may need quite a few sections to collect all of the in-focus light in that sample to render your 3D volume.
So the way that I would advise people to set up confocal volume imaging is to set your microscope to start collecting the Z slices right at the top of your sample, and you should have no light in-focus at that point. So the specimen should be just out-of-focus at the top, and then it should move through all the way down to the bottom where there's no in-focus light at the bottom either. If you're going to try and collect the perfect 3D section, you'll need to scan with the optical slice optimized, and that may take some time.
So that's really the ideal way of collecting your 3D sections. Now, I'm just going to move on and talk about some of the common questions I get asked in my core facility, when people start using confocal microscopy. The first question I get asked by people who've never done any confocal before, is when I look down the microscope and it's set to scan on the confocal, I can't see anything. There's good reason for that, because the laser light is so powerful it would damage your eyes if you looked at it. So if you need to see your samples down the microscope, you need to ensure that the light part is set to your eyes. If it is and you can't see anything, then it may well be that your sample staining hasn't worked. When you set the light to confocal mode, if you can't see anything then that's a good thing, because that means that the lasers aren't actually going to go to your eyes, and if you can see something then you need to contact your microscope manufacturer immediately.
The second question is: While I'm imaging, my sample fades away. That's to do with the fact that the lasers on the confocal are set to very high power, and so they can sometimes bleach your dye that you're using. So the best way of avoiding that, there's nothing you can do once your sample is bleached, the staining has gone, but maybe in a different place on the same slide, you might have another opportunity to acquire some images. The thing to do there is make sure that the detectors are set to maximum sensitivity, and the lasers are on the minimum power possible, that way you will minimize the bleaching in your sample.
The next concern people often have is that their sample looks really dim. Now, if your sample looks really dim by eye, then the staining is weak and the confocal will need optimizing. So you may need to have your detectors on maximum sensitivity, and use a laser a little bit more powerfully, even possible to just get any sort of image. Although you do need to remember that you need to be representing your data fairly, and so producing something that's dim on the slide is probably the right thing to do.
If the staining looks really bright by eye, and you need to be honest about this, the detectors may need to be made more sensitive because maybe it's just been left by the previous user in a space where they had great staining, and your staining is not the same as theirs, so you'll need to re-optimize the machine. But I will reiterate the point that it's very important to take a representative image of your sample.
If you're on a confocal and your sample looks really grainy, that might be because a confocal is still in its initial fast scan mode, which uses for you to optimize the detectors and make sure that everything is within the linear range of the detector, so it's not saturated or undersampled. What you can do is you could actually choose to slow down the scan speed on your confocal, and also increase the averaging of the frames. So this means that by slowing down and letting the laser dwell on each pixel for a little longer, and by averaging maybe two or four frames, you'll cut down this grainy noisiness and you'll really get a nice looking image.
The last question's a tricky one. You think you see co-localization in your sample, but your mentor, your PI or your group leader, your postdoc, or whoever doesn't agree with you. So if you just show one image of the co-localization, then it's not quite enough really to make a robust scientific case. What you need to have when you're presenting co-localized data, is all of the controls present for your staining, and you need to make sure that you're capturing your images using the same settings on a confocal that you use for the controls. So you need to have an image of just the green on its own, and then nothing in the red channel; the red channel on its own and nothing in the green channel. It's also very important to set the instrument up using sequential scanning.
This is the end of the bit of the webinar where I'm talking about theory, and I'm just going to summarize what I've said. So confocal microscopes are there to remove out-of-focus light, and produce high-quality images. Confocal microscopes mostly work by point scanning, and they will scan each image point-by-point, and the speed and the averaging on your confocal can be altered to improve your image quality. Sequential scanning is important to stop one dye bleeding through into another channel, and that's very important when you're setting up a co-localization experiment. Confocals can be used to reconstruct 3D volumes, but it's very important to understand how thick your optical section is, and a little bit more information about the objective and its numeric aperture at the pinhole when you're setting up your 3D volumes.
So now I'm going to hand over to Miriam to give a summary of the products and services which are available.
MF: Thank you very much, Ann, for such an informative and detailed seminar. Hello again, everyone, I would like to take this opportunity to tell you a bit about Abcam microscopy resources and products that can help you to improve your cell imaging experiments. Rabbit monoclonals or RabMAbs offer high affinity and specificity, which results in higher sensitivity and low background staining. This makes them ideal affinity reagents for demanding applications, such as IHC or formalin, paraffin-embedded tissues. To give you the best experience, each RabMAb has been tested on multiple human tissue arrays for IHC, and on multiple samples for ICC/IF. RabMAbs also offer diverse epitope recognition of human plotting targets, and the mouse orthodox, so there is no need to generate a separate surrogated antibody. Because they are rabbit generated they are ideal for use on mouse or rat tissue samples. They can also be easily paired with mouse or rat monoclonal antibodies for simultaneous staining. For further information please visit abcam.com/RabMAbs.
For staining with your RabMABs we recommend one of our Alexa Fluor® conjugated secondary antibodies. We have just launched our new range of Alexa Fluor® conjugates 405, 568 and 750. All the secondary antibodies have been extensively tested in the Abcam laboratories, in order to guarantee bright staining and low background. The selection of pre-adsorbed antibodies is large, and ensures low species cross-reactivity. The dilution range for all these products is between 1/200 and 1/1,000, and the products are very competitively priced. To find out more, go to abcam.com/Alexa.
The Abcam catalogue also includes a whole range of cell imaging tools for multicolor staining. Discover our CytoPainter range of kits and reagents for staining actin filaments, ER, Golgi, mitochondria and lysosomes in multiple colors. It is an easy way to study co-localization without having to fiddle around with multiple antibodies. CytoPainter products can be used in combination with secondary antibodies and nuclear dyes. The image on the top right corner shows you an example of multicolor staining of mouse embryonic bodies.
If you want trouble-free staining of nuclei, why not try our far red dyes that will display nuclear staining in just 5 minutes. Included in this range are DRAQ5 and DRAQ7, which can be used for staining live or fixed cells, respectively. Here, in the bottom image, you can see a nuclear staining with DRAQ7, while the blue staining originates from an AMCA conjugated secondary antibody.
Our extensive portfolio of validated secondary antibodies for cell imaging includes pre-adsorbed Alexa and DyLight conjugated secondary antibodies for minimal species cross-reactivity. Our range also includes chromeo conjugated secondary antibodies for STED microscopy and AbGold conjugated secondary antibodies for high-resolution electromicroscopy. To increase tissue penetration in IHC experiments, we recommend you to try one of our F(ab')2 fragment antibodies. For non-fluorescent imaging or enzymatic detection, Abcam also offers a comprehensive range of products for immunohistochemistry. Included in the portfolio are EXPOSE-IHC kits which provide greater sensitivity in comparison to polymer and ABC detection systems to a smaller detection complex. The IHC portfolio also includes the classical biotin and streptavidin kits, and ancillary reagents. If you would like to know more about all this range of cell imaging products, please visit abcam.com/imaging where you can find more information.
As Ann has mentioned during the webinar, it is very important to protect your slides from photobleaching. For this specific purpose, Abcam also has a range of fluoroshield mounting reagents. These are available with or without nuclear dyes, such as DAPI and PI.
Abcam scientific support team is here to answer any questions you may have. The team members are multilingual and offer support in a range of languages, including French, Spanish, German, Chinese and Japanese. You can contact them in the US, UK, Hong Kong and Japan on the email addresses and phone numbers shown here.
On the Abcam website you can find a variety of free resources and protocols. Of potential interest to you are our brand new “Multicolor imaging tools for cellular staining” poster, and the “Understanding secondary antibodies guide”. The latter explains when to use F(ab')2 fragments and pre-adsorbed secondary antibodies in multiple staining experiments. Please contact us at email@example.com for your free hardcopy, or download a PDF at abcam.com/posters.
You can also listen to all our past webinars, including other microscopy-related ones, such as “How to design multicolor staining” and “How to design IHC/ICC experiments” by simply going to abcam.com/webinars.
As a thank you for attending this webinar, we would like to give you a special 25% discount on all secondary antibodies, CytoPainter products, IHC kits, avidin/streptavidin kits and RabMAbs. All you have to do to take advantage of this offer is to quote the promotion code that appears on this slide when you're placing your order.
Now, the part you've all been waiting for, the answers to the quiz. The correct answers are shown in bold, so for question one: Which fluorochrome is not typically used in microscopy? The answer is PE or phycoerythrin. This dye exists in vitro as a 240 kDA protein with 23 phycoerythrobilin chromophores per molecule. PE is the brightest fluorochrome for flow cytometry, but since it's highly photobleachable it is unsuitable for fluorescent microscopy.
For question two: Which color combination will most likely give you the spectral overlap? The answer is A, mainly because the fluorochromes Alexa 555 and 648 have quite a high spectral overlap.
For question three: Which of the below fluorochromes have the highest quantum yields? The correct answer is Alexa 488, as it has a quantum yield of 0.92, so it's very close to one.
I would like to finish by thanking Ann again for such an interesting webinar, and all of you for attending. Just to remind you that when you log-out from the webinar you will be redirected to a page where you can download the slides, and where you can find the promotion code and terms and conditions. So, now, Ann is happy to answer any of your questions.
AW: Thank you very much, Miriam. I've got a few minutes to answer some of the questions that you've all kindly emailed in. So the first question I'd like to answer are: What are the criteria for a good quality image from confocal microscopy for publishing? Which is an excellent question. The first thing that's probably the most important is it needs to be very representative of your sample, so you need to make sure that maybe not only just you, but you and somebody else; maybe your supervisor or a colleague in your lab agree that what you see down the microscope is what is also being seen by the image that is produced on the confocal.
Something that you will need to know when you're trying to publish your image at publication quality, is that it needs to be at a high-resolution, it needs to be about 300 dpi otherwise it will look grainy and nasty when you actually upload it onto the website for the publication. It's quite important - these images will be put in your paper, and they'll come out a lot smaller than what you see on a confocal screen. So it's quite important to really make sure you've got that polishing there. You need to make sure the background is properly black.
If you can, try and exclude any artefacts in your image, so it might be possible for you to turn around the area that you're scanning on the confocal a little, so that you can exclude anything you don't want; if you've got a nasty blob of rubbish that's unfortunately ended up next to your perfect cell or tissue.
Do make sure that everything is within the linear range of your detector, make sure your background is black, but it is linear under the detector and nothing is saturated, and try and improve the focus as much as possible. So you would want to slow down the scanning speed, and also include the averaging for a publication quality image.
The next question is: How can you tell the difference between autofluorescence and weak staining? That's a great question, that's a very difficult one to answer, and it's posed quite a lot of problems I think for people. The best way of doing that is actually by just having control where there is nothing stained, and so you just have your tissue or your cells.
All confocals that I've ever worked with have got a brightfield option, and so what you can do is you can focus your sample by brightfield, and you can even get the confocal to acquire a brightfield image for you generally. Then you can just acquire an image of the sample in brightfield, so it's in-focus and then having the settings for the confocal set up for acquiring whichever dye it is you were going to acquire, so Alexa Fluor® 488, just take the image with nothing there and see how the autofluorescence looks. What you then need to do is obviously use those settings and if you can see nothing, you have to optimize them, and so the autofluorescence is minimized, and then you can actually take your image.
There's advanced tools called spectral and mixing which is it's possible to do with certain confocals. I'll just mention the concept of spectral and mixing here, I won't go into it in detail because it's an advanced application. However, if you can look on the internet for this, there's plenty of resources available for it online. It may be possible that if you're fortunate enough to work in the manager facility, that your facility manager may know about spectral and mixing. Usually, the emission spectra for the autofluorescence is slightly different from the emission spectra from the dye that you're using. So you can collect the emission spectra from the autofluorescence and remove that, but that can be a little bit more demanding to do.
But it is certainly possible to differentiate between autofluorescence and weak staining, and what I'd try and recommend is you optimize your staining as much as you possibly can to try and get it a bit brighter, if possible.
So I just have a question from somebody asking: What is a perimeter pixel dwell? So if we go back to the slide in the presentation and when we're talking about your image. What happens is, well, the way the confocal sees this is a series of little boxes and the boxes are called pixels, and the laser is moving across the image pixel-by-pixel. Pixel dwell is basically the length of time that the laser is sitting on a given pixel before it moves. So if it's sitting there for a millisecond or a microsecond that will depend on how the image looks. Usually, the longer the pixel sits on a given point so the longer the dwell time, the less the noise is, but the more chance there is of your sample being bleached.
Somebody else has asked: Can I do a deconvolution of my images on the confocal microscope? Certainly you can, and for some applications it's worthwhile bothering. So if you're really pushing the limits of the resolution of what your confocal can do; so if you're trying to image something that's about 200 nm and you're trying to do a 3D volume of this. For instance, for imaging the nucleus then you may want to try and do some deconvolution. Deconvolution, in principle, is there to remove out-of-focus light. A confocal pinhole, in practice, is also there to remove out-of-focus light, so really you only need to deconvolve your images if you're a bit uncertain if there is any out-of-focus light remaining in your confocal images when you're done with your confocal microscopy. Some people choose to do this, others don't. I generally advise not to do it unless it's something very, very specific and detailed, and it's right at the limit of resolution of a confocal. But I'd say that's something that's really down to you, your group leader and others in your group as to how things happen in your field.
I think we may have time for a couple more questions. So I have a question saying: Do all dyes emit at the same wavelength as they're excited with? Well, so fluorescence, the answer to that question is, no, they don't. So chromogenic dyes, which are the sort that are used for histology, like H&E, generally speaking, they're just little chemicals and they emit light if you shine normal light at them and they will be a color, and you'll be able to see that. Fluorescent dyes aren't like that, they're different, and so, basically, the fluorescent dye is excited by a high-energy light source, so it absorbs high-energy photons and then it has to lose some of the energy because of the way the dye is structured, and then it needs to emit light at a lower wavelength. So fluorescent dyes emit light at different wavelengths from the light that they're excited from, and the wavelengths that they emit at depends on the chemistry of the dye. So you need to go back and check out what the absorption or excitation spectra for your dye is, and then what its emission spectrum is before you can set things up.
Somebody else has asked: Please can you explain the order of use of the color channels in confocal? Well, to be honest, I would say that depends on the instrument that you're using. Usually, people will collect the shortest wavelength first, so they'll collect the blue channel first and then the green and the red. But, really, what happens is when you're simultaneously scanning they are all collected all at the same time, and then the light is split up by the dichroic mirrors which are in the emission light path. So the dichroic light will send some wavelengths in one direction, and other wavelengths in other directions. If you're using sequential staining, then basically it will collect blue, green or red, whichever order that you put it in, and it's kind of up to you, really. The confocal will be pre-set for certain channels and if it's easier when you're starting to use a confocal to start off with, then just use the pre-sets, that will be the best advice that I have.
So the last question is: Can confocal be used for formalin or/and paraffin-embedded tissue, or just frozen tissue? The answer to that question is it can be used for both. For formalin and paraffin-embedded tissue it's a little bit more tricky to do the set up, you need to do some sort of antigen retrieval step. There are protocols available for this, I think this may be covered in other Abcam webinars, so I wouldn't be able to advise in detail about this now. But it certainly is possible to do that, you just need to be quite careful about how you set up that for confocal microscopy, but either sample will work and we have published that.
Well, thank you very much for all of your questions, and thank you for your time. I'll just hand you back to Lucy now.
LP: Thank you very much, Ann, and thank you to yourself and Miriam for presenting today. Unfortunately, due to time restrictions we have not been able to answer all the questions received. For those whose questions were not answered, our scientific support team will be contacting you shortly with a response. The webinar presentation just given is available for download. When you log-off from the webinar you will automatically be redirected to a webpage where a PDF of the presentation can be found. If you have any questions about what we have discussed today or have a technical enquiry, our scientific support team are always very happy to help and they can be contacted at firstname.lastname@example.org. We hope you have found this webinar informative and useful to your work, and we look forward to welcoming you to another webinar in the future. Thank you again for attending, and good luck with your research.
Ladies and gentlemen, that does conclude today's webinar. We'd like to thank you once again for your time today, and you may now disconnect.