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| Frequently Asked Questions | Manuals | |||||
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| CSIF General Policies |
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| Q. What is the mission of the CSIF? |
A. Current microscopy technology is getting more and more complex and requires a greater investment in money, time to learn how to effectively operate the equipment and in maintenance. The CSIF was established as a core facility to provide access to high end systems that would make these technologies more accessible to casual users. Our functions are:
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Q. Who is allowed to use the CSIF? | A. The Cell Sciences Imaging Facility operates primarily for Stanford University faculty, post-doctoral fellows, graduate students and staff. A limited amount of time is available for people from affiliated institutions and local biotech industry. |
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Q. How can I schedule time on equipment? | A. We have an online reservation system for trained users to reserve time on the individual microscopes and computers. It can be found at http://taltos.stanford.edu/schedule. You will need to register to use this reservation page, so new users must contact CSIF staff to arrange training prior to using the scheduling software. | |||||||||||||||||||||||||||||||||||
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Q. Why doesn't the scheduling software allow me to delete my reservation? | A. If the time you are trying to delete is less than 24 hours in the future (i.e. the day of reservation), then the software will not allow the deletion to be made. CSIF policy requires 4 hours advance notice of a deletion to allow other users ample time to book unwanted time slots. To cancel and not be charged for reserved time, on the day of reservation please contact the CSIF staff and they will assist you. | |||||||||||||||||||||||||||||||||||
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Q. How do I arrange a training session? |
A. Due to the complexity of the equipment, we do all training sessions on a one-on-one basis. To arrange a session, contact CSIF staff at 3-2449 to schedule a mutually convenient time. You will need to provide a sample stained with the fluorophore(s) you will be using in your experiments. It is advisable to wait to schedule time until you have worked out an effective staining protocol. | |||||||||||||||||||||||||||||||||||
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Q. What should I bring to training session? |
A.You will need a WELL LABELLED specimen. That means that you must practice your staining protocol and look at your samples prior to coming to the training session. Verify that the signal is strong (easy to see) and that the background is nearly black. All of our systems are based on inverted microscopes, so the sample must either be:
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Q. How are labs charged for services? | A. The CSIF accepts SU-13 (interdepartmental transfer) forms signed by the proper departmental official or email authorization from the departmental administrative official. In both cases, we require a valid account number (PTA), the lab PI's name (and lab members who can use the SU-13, if appropriate) and the months the SU-13 is valid for. Account authorization may be specified to cover any number of months within the current fiscal year (Sept 200x to Aug 200y). Because of the independent use of users and monthly billing policies, we cannot honor "not to exceed" requests, but will close the SU-13 in the month when billing first exceeds the dollar amount specified. | |||||||||||||||||||||||||||||||||||
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Q. How much space does each user get on the server and how long is it stored? | A.
All users:
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| Confocal Laser Scanning Microscopy |
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Q. What is confocal laser scanning microscopy? | A. In a conventional epifluorescence microscope, light of a defined wavelength is reflected by a chromatic reflector through the objective and bathes the whole of the specimen in fairly uniform illumination. The chromatic reflector has the property of reflecting short wavelength light and transmitting longer wavelength light. Emitted light from the specimen which is of longer wavelength passes straight through the chromatic reflector to the eyepiece where it is detected by eye. The major problem with conventional microscopy is that up to 70% of the collected light is from out-of-focus interference. In thick specimens, this commonly results in an inability to discriminate the location of labelled proteins.
Confocal laser scanning microscopy (CLSM) was developed to overcome the problems associated with out-of-focus light. In a CLSM system, a single point of excitation light is scanned across the specimen. The point is a diffraction limited spot on the specimen and is usually produced by focusing a parallel laser beam. With only a single point illuminated, the illumination intensity rapidly falls off above and below the plane of focus as the beam converges and diverges, thus reducing excitation of fluorescence for interfering objects situated out of the focal plane being examined. Fluorescent signal passes back through the dichroic reflector and then passes through a pinhole aperture situated in a conjugate focal plane to the specimen. The pinhole acts to restrict light passing to the detector to that emitted from the focal plane. Light rays from below the focal plane come to a focus before reaching the detector pinhole, and then they diverge so that most of the rays are physically blocked from reaching the detector by the leaves of the detector pinhole. In the same way, light reflected from above the focal plane focuses behind the detector pinhole, so that most of that light hits the leaves of the pinhole and is not detected. Light passing through the image pinhole is detected by a photomultiplier tube. In summary, a confocal imaging system achieves out-of-focus rejection by two strategies: a) by illuminating a single point of the specimen at any one time with a focussed beam, so that illumination intensity drops off rapidly above and below the plane of focus and b) by the use of a pinhole aperture in a conjugate focal plane to the specimen so that light emitted away from the point in the specimen being illuminated is blocked from reaching the detector. |
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Q. Does CLSM bleach samples faster than conventional immunofluorescent microscopy? | A. Fading/bleaching of a labelled specimen is a major problem in fluorescence microscopy. The higher power and focussed beam of a LSCM causes enhanced fading of a specimen as compared to a conventional epifluorescent microscope which essentially bathes the entire specimen in lower power, wide-beam excitatory light. However, this is balanced by the shorter duration time that the laser is illuminating the specimen in a LSCM as only a tiny portion of the field is illuminated at any one time in the scanning process. | |||||||||||||||||||||||||||||||||||
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Q. How do I clean the confocal microscope at the end of my session? | A.Take a piece of lens paper and squirt a spot of Windex in the center. Place the lens paper on the objective on the wet section, pull the flat lens paper across the objective, moving to a clean (dry) section of the lens paper. Do NOT scrunch up the lens paper and scrub the objective as this may scratch the objective. Use Kim Wipes to clean the stage and any other parts of the microscope that might have oil on them. A neat and tidy environment makes everyone happy (especially the CSIF staff) and gives you a better chance of getting good pictures. | |||||||||||||||||||||||||||||||||||
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Q. I have trouble finding my sample when I look in the eyepieces. Are there any special tricks that I can use to find it more easily? | A.Check the following:
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Q. Why can't I focus on my specimen? | A.The high magnification objectives on the microscope all have fairly high numerical apertures (~1.4). This means that the depth of focus (how far into the sample you can focus) is very small, so your sample must be close to the objective. The most common problems are:
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| 2-Photon Scanning Microscopy |
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Q. What is 2-photon scanning microscopy? |
A.The two photon microscope depends on the 2-photon effect, by which a chromophore is excited not by a single photon of visible light, but by two lower-energy (infrared) photons that are absorbed contemporaneously (within 1 femtosecond). Fluorescently-labeled specimens are illuminated by a titanium:sapphire laser that produces very short (less than 200 fs) pulses of infrared light--with a very large peak amplitude (50 kW)--at a rate of 76 MHz. Fluorescence from the two-photon effect depends on the square of the incident light intensity, which in turn decreases approximately as the square of the distance from the focus. Because of this highly nonlinear (~fourth power) behavior, only those dye molecules very near the focus of the beam are excited. The tissue above and below the plane of focus is merely subjected to infrared light that causes neither photobleaching nor phototoxicity. Although the peak amplitude of the IR pulses is large, the mean power of the beam is only a few tens of milliwatts, not enough to cause substantial heating of the specimen. Or to answer another way: The traditional drawback to using fluorescent markers in conventional microscopy is that out-of-focus information is collected, making the precise localization of proteins within the cell or tissue difficult to achieve. Current solutions to this problem include confocal laser scanning microscopy or wide-field deconvolution technologies to generate optical slices that include only in-focu information. | |||||||||||||||||||||||||||||||||||
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Q. What are the benefits of 2P excitation? | A.
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Q. How does 2 photon microscopy differ from confocal laser scanning microscopy? | A. Problems with normal confocal fluorescence microscopy:
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| Wide-Field Deconvolution |
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Q. What is deconvolution and how does it work? |
A.The convolution and deconvolution concepts have very precise mathematical definitions. If you're interested in these, please find a textbook and read further about it there. What we're trying to do here is to give a more conceptual idea of what deconvolution means in a microscopy context. A limitation to fluorescence images captured by a light microscope is that out-of-focus fluorescence can obscure data in the plane of interest. In laser confocal microscopy, this problem is eliminated by only collecting light generated in a specified plane. The drawback is that the fluorescent dye is excited by a high intensity laser. This leads to photobleaching and phototoxicity. Deconvolution provides a computational solution to this problem by calculating and subsequently subtracting the component of a fluorescence image that comes from planes above and below the plane of interest. A point spread function (PSF) is determined for each objective by measuring a fluorescent bead, which mathematically describes how fluorescent light from a single point will be diffracted. During deconvolution the Fourier transform of the PSF is applied to the raw data images to reassign the out of focus information back to the point where it was emitted. The result is a deconvolved image in which details can be distinguished more clearly. |
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Q. What are the different kinds of deconvolution? | A.
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Q. Is blind deconvolution better than calculated PSFs for cells due to varying refractive indexes in different parts of the cell? | A.The following is a question from the confocal newsgroup: If I look at a cell in embedding medium, Ri (refractive index) is likely to be different in the medium and the cytoplasm and maybe even between cytoplasm and nucleoplasm. All or at least 2 Ri's will be different from the Ri of the immersion medium. I understand that the PSF changes dependent on the Ri. So, in theory, it might be a good idea to use different PSFs for different areas of one image stack (disregarding the necessary computer power). I guess this would be quite difficult to realize with measured or calculated PSFs but I don't see a principal reason why this shouldn't work with blind deconvolution. Is this done or does blind deconvolution as implemented these days use only one PSF per image? In the latter case, the once determined PSF could as well be used for other 3D-stacks recorded from the same slide on the same day under the same conditions, probably cutting down calculation time considerably. The Answer: Yes, the basic pre-requisites are given with blind methods. However, consider that a true space variant method requires the same computational effort that is used now for the whole image stack times all the discretisation units. Even if you neglect aberrations in transverse dimension you still have N-slices to get object and PSF distribution from. An efficient way might be the blind deconvolution of small blocks. They would have to be as large as to hold the 3D PSF region of support. This block is then shifted through the entire volume and the estimation of PSF and image takes place in each one. Well, I would understand if that has not become a commercial product yet. The effort is astronomical! The blocks might also be shifted in larger steps, e.g. that they become adjacent, that would greatly reduce that effort but then you have to consider each block as the result of a space invariant restoration with a different PSF for each. That might be a first step considering todays computers. |
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Q. What are the nitty-gritty details of deconvolution? Jason Swedlow speaks from experience. |
A.Excerpted from the confocal newsgroup: The problem with deconvolution microscopy is that this term refers to many fundamentally different approaches and the current nomenclature doesn't clearly distinguish between them. You have to be careful which you choose. To my mind, the major difference between the methods listed above is that the neighbor-based methods radically change intensity relationships such that subsequent quantitative analysis, ratioing, etc. is not possible. In general, the various iterative methods preserve the intensity relationships, so you can compare the amount of nuclear and cytoplasmic staining, etc. So it depends what you need to use the images for. Regarding the use of empirical vs. calculated PSFs, if possible, it would be "better" to use the empirical PSF. This makes intuitive sense, and the comparison has been done between empirical and theoretical PSFs, but it hasn't really been carefully shown for empirical vs blind deconvolution on samples with a wide range of S:N. And no, it's not enough just to show that the image looks "better" - a true test requires known standards (but a standard can be real, like a microtubule), a range of S:N levels and preferably a frequency-domain analysis of the restoration. Tim Holmes' group has presented these types of analyses and shown that blind deconvolution works very well in their images. Moreover, there is a version of AutoDeblur especially designed for LSCM data. While any deconvolution method can theoretically deal with LSCM data, AutoQuant has really pushed the development of specific filtering strategies to handle LSCM data. Then how do the iterative methods differ? I'll quote from my own experience - not by much, if you just consider the algorithms themselves. That is, they all restore the types of images people use to demonstrate the methods (typically good S:N) and give significant improvements in contrast and S:N. The major differences are implementation. Specifically, what types of filtering are used, whether the whole image is handled or parts divided up, how long it takes to run, type of PSF, etc. There are published comparisons between different methods, but usually one group has rewritten another group's method and omitted all the "tricks" (filtering, padding, etc.) that make these things work. Yes, it would be nice to have a clear comparison, but we would still argue about what types of samples to use. A true comparison will be hard now, since most of the code is in the hands of commercial vendors. As this thread shows, all the products have customers that swear by their results. I'm tempted to leave it at that. But what about that pesky empirical PSF? Is it that hard to measure?? Well, your first one won't be right. But as others have said, it's a matter of knowing what you're doing, i.e., having a good protocol (just like running a gel!). But a "bad" PSF is telling you something. Any of the problems that occur during a PSF measurement (e.g., stage drift, refractive mismatches in the immersion media, temperature changes due to heavy ventilation, lamp flicker, camera noise, etc.) also occur during your experiment (the "biology" that we all want to do). In short, the PSF is reporting on the errors in your imaging. You can choose to ignore it, and deconvolve using a theoretical PSF or use blind deconvolution. But all those errors are still there and blind or any other type of deconvolution, because of the assumptions made in the PSF, can't do anything about it. (In principle, it would be possible to deal with stage drift, scattering, etc. in software, but there are no commercial versions of these corrections). Alternatively, you can use the errors in the PSF measurement to suggest ways to improve your microscopy. Then you can decide what deconvolution method you want to use (blind, empirical, etc.). And just reading the thread on this one shows that the choice involves performance, as well as cost, ease of use, etc. A VERY IMPORTANT POINT: To my knowledge, all commercially available iterative deconvolution methods assume a radially and axially symmetric PSF (the averaging is done in Fourier space in all cases, I think). The axial symmetry is most dependent on the amount of spherical aberration present in your image. WHEN USING IMMERSION OBJECTIVES, IT IS SO HARD TO SET UP A NON-ABERRATED IMAGING PATH THAT YOU SHOULD ASSUME YOUR IMAGING LIGHT PATH GENERATES SPHERICAL ABERRATION. THE DECONVOLVERS HAVE BEEN SCREAMING ABOUT THIS FOR SOME TIME; IN THE LAST FEW YEARS EVEN THE CONFOCAL TYPES HAVE PICKED UP THE CALL. It's easy to see - just look for asymmetry in the out-of-focus rings above and below a bright fluorescent source in your sample. If you see any intensity asymmetry (i.e., brighter above the object than below) as you focus up and down, you have spherical aberration. An image that contains spherical aberration has degraded axial resolution and decreased signal (same signal spread out over a larger volume). Regardless of the deconvolution method you use, YOU WILL NOT CORRECT FOR THIS. What to do?? You'll need to 1) adjust the refractive index of the immersion medium or 2) get a lens with a correction collar (like the 60x/1.2 water immersion lenses that have become available over the last few years). This sounds hard, but once you've picked the right oil (or the right setting of the correction collar) for your sample it's over with. THE POINT: If you ignore what the measured PSF is telling you, you will record an image with degraded signal-to-noise and resolution. Why should you care?? Your major interest is the biology?? Simply because when you go to determine what the image is telling you, (e.g., whether two or more components are colocalised), the aberrations present in your image will give you false results - your "biology" will be wrong. And no amount or type of deconvolution or LSCM will change this. BUT THIS TAKES TIME AWAY FROM ME DOING AN EXPERIMENT!!! Yep, but in general you have to do the experiment correctly. If you can get away without worrying about some of the details, then do it. But be aware of the limitations. And wouldn't it be better not to have so many caveats? And finally: BOY IT TAKES A LONG TIME TO DECONVOLVE!!!! This varies widely depending on the actual hardware and software. As I said, on the software side, there are major differences in implementation (as well as type of methodÑnote there are five different algorithms in the AutoQuant package. Deconvolution calculations require a number of large arrays representing different forms of the image to be stored simultaneously. This means that large amounts of data must be moved around inside the machine, so computers with very fast buses actually perform much better even with nominally slower processors (the clock speed actually isn't the best performance spec anyway). If you have the means, SGIs are built to move large amounts of data quickly, so they really perform in these applications (nope, I don't own SGI stock). Scared of Unix?? We have a number of non-specialists using our SGIs and they don't seem to mindÑ all the file management etc looks like Windows or Mac. Plus, unlike the Windows or Mac boxes, they are truly multi-tasking, so they don't crash nearly as much. Our current spec: 10 iterations of constrained iterative deconvolution on a 512 x 512 x 64 image takes about 3.5 minutes on a dual processor R12000 Octane. This beats the real data collection time (find the cell, set the imaging conditions, take the image), so the experiment is now limiting. |
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Q. What excitation and emission filters are installed in the API DeltaVision microscope? |
A.All of the filters on the DeltaVision are band-pass filters. The following table provides the filter name (for the most common fluorophore that the set is used for), the excitation filter (center wavelength/width of band, in nm), and emission filter (center wavelength/width of band, in nm).
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Q. What numerical apertures (NA) and working distances of the objectives on the DeltaVision? | A.The Deltavision deconvolution microscope has 5 different objectives with the following NAs:
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Q. What is the lateral resolution of the DV and the confocal scopes? | A.Technically, resolution is limited by the resolution of light and the optics of objectives. For most samples, it is approximately 200 nm. In a confocal system, this is true for the xy resolution. The axial (or z) resolution in a confocal is only 1/3 of this (about 600 nm). The deconvolution algorithm partially gets beyond these limits and operates at slightly higher resolution (Applied Precision will say that the resolution of the DeltaVision is about 150 nm both xy and z under optimal conditions.) |
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Q. Why can't I see anything in the eyepieces on the DeltaVision microscope? |
A. Check the following:
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Q. Why is first image collected different than what was seen when I determined the bottom of the sample? | A. This is a common problem and is due to a number of factors. The motor has moved the objective back to the same physical place, but if your sample moved, then what you see is different. Common reasons for sample movement: 1 - The slide moved on the stage Correction: Clamp edges of slide before focussing 2 - Too much or too little immersion oil used, causing tension or vacuum between objective and coverslip Correction - clean both slide and objective and apply small drop of immersion oil on objective 3 - Sample is not fixed to coverslip and movement of objective results in movement of sample Correction – Put sample on stage and wait fo sample to settle. Calculate top and bottom of specimen as outlined in step 1b, and leave objective at bottom. Wait about 30 seconds, acquire image. If sample doesn’t move – then collect series. If sample moves, then you need to modify sample preparation to stabilize your sample. |
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Q. How much out of focus should I be when setting my endpoints for collecting a stack of images? |
A. This is a hard question to answer. The reason you have to be somewhat out of focus is that the deconvolution algorithm uses out of focus information to mathematically filter away light originating from points that are out of focus. Normally, if you move in 1 micron increments, then stop at the first section in which the entire image is just slightly fuzzy (therefore, about 1 micron past the last "in focus" image). |
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Q. I get the "camera saturated" message. What should I do? | A. The camera gets saturated when too much light reaches it. There are two ways to reduce the amount of light that reaches the camera:
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Q. What is binning and when should I use it? | A. The camera chip can record a field that is 1024 x 1024 pixels. Usually, if 1024 x 1024 pixels are collected, then all 1+ million pixels will be displayed. However, when samples have a low fluorescence, then the intensity recorded for each pixel may be very low. One way to increase the intensity per pixel is to make each pixel refer to a larger area of the sample. This is effectively zooming OUT. On a system such as the DeltaVision, this is accomplished by collecting 1024x1024 pixels and then ADDING up each 2x2 set of pixels (so 4 pixels total) and the resulting image is now a 512x512 field. Think of it as putting 4 pixel intensities into a single "bin." The example above is a 512 field with 2x binning. There are other levels of binning. The math is that 1024/output field size in pixels = maximum bin factor. (So 256 can be binned 2x or 3x, while 512 can only be binned 2x, etc.) There are 2 sets of circumstances where this is useful 1) staining is very light, such that very long exposure times must be used to get reasonable dynamic range of measured intensities (max – min > 400) or 2) you want to see a larger field size, but don’t want to switch back and forth between high power (oil) and low power (air) objectives. |
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Q. The deconvolved dataset doesn't have the deltaV icon, how can I open it? | A.This is a bug in the software which happens only on the SGI Octane architecture (memnoch). There are two ways of solving this problem
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Q. The SoftWoRx program just froze. What should I do? | A.Don't panic. This happens once every now and then. If you can still move the cursor, there are a few things you can try
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Q. Is there any way to make the deconvolution go faster? |
A.The deconvolution time is dependent on the amount of data you wish to deconvolve and what else the CPU is being called on to do. Therefore, a number of possible ways to decrease the deconvolution time are:
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Q. Why doesn't deconvolution improve the quality of my images? |
A. 1) Make sure that you have selected an exposure time that uses the full CCD array detection range. The camera can register values between 0 and 4095, and if you only have a difference of 200-400 between your min and max values, this means that you are only using 5-10% of the available detection range. The reason your images still look fine on the screen while you collect them is that you're looking at a scaled version of them. The display can only show 8 bits, i.e. 256 different values, so whatever range you've captured is scaled to 0-256. 2) If your output intensity values are over a wide range, check the final r value for the deconvolution. Ideally, at iteration 15 the number should be 0.1 - 0.3. If your values are between 0.3 - 0.5, you most likely have a problem with your sample preparation which led to spherical aberration (the cover slip thickness, amount of mounting media, refractive index mismatch) and which can probably be corrected for by slight changes in your sample preparation protocol. If the r value is greater than 0.5, then there is a larger problem - difficult sample type, too low intensity of stain, etc. |
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Q. How do I deconvolve only a portion of a stack of images? |
A.When you choose Process - Deconvolve from the SoftWoRx toolchest, a window opens up which prompts for the raw dataset you wish to deconvolve. Click the "Details" button and enter the section numbers corresponding to desired subset of sections (in the Z start/end boxes). Click the Close button. Now the deconolution will only be performed on the z-sections you've chosen. |
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Q. How do I make an animated volume rendering? |
A.In the SoftWoRx window, click the View menu and choose VolumeViewer. This will open a new window in which you can select a number of parameters for the dataset that you wish to render. Under "Viewing Parameters," you will choose the algorithm to generate the volume view. The three options we recommend are Additive, Max Intensity, and Progressive. Additive displays the total of all intensity values for each pixel along your "line of sight". Max Intensity displays the highest intensity value of the pixels along your "line of sight" and thus reduces the background compared to the Additive algorithm in most cases. Progressive should be used when you wish to generate a surface view of your image stack. Other parameters you will need to select are the z resolution, the quality, and what type of projections you want to make. All of these are pretty self-explanatory and are chosen depending on the volume view you desire. If you have any questions about these, please ask the CSIF staff. |
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Q. How can I save the scale bar information when I export files to TIFF from SoftWoRx? |
A.To save scale bar information, you use a SGI utility called Media Recorder (formerly Image Snapshot in older versions of IRIX - pre-6.5) to capture the screen output (which will include the scale bar). In the SoftWoRx menu, select Utilities - Image Snapshot. This will open up a "Media Recorder" window and a marching ants marquis. In the Media Recorder window, to go Tasks - Image - Custom Settings and under File Format choose TIFF. Then move the marching ant boundaries until they enclose the edges of the image you want to save (move cursor to a line of marching ants - double headed arrow will appear, grab arrow and move to desired position). Hit "Record" (button with red filled circle). This will save the output in your home directory (/usr/people/username) as "imageX.tif) where X is a number that increments by one every time you save another picture. You can change the name by opening your directory and clicking on the filename and retyping it to the name you want. Make sure you hit return when you are finished typing (otherwise the name reverts to the original name). When you hit record, the marquis disappears. If you want to do a second screen snapshot, go to Edit - Show Selection Frame and it will reappear. |
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Q. How do I save only a portion of a stack as TIFF images? |
A. Open the dataset you wish to save. Choose "Save as TIFF" under Files in the Image window. Click the "Details" button and enter your starting section # and ending section # as the z start and z end points. Click the "Close" button. When you hit "Do It" in the main "Save as TIFF" window, only the sections you've specified will be saved as TIFF files. |
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Q. What bit depth (8, 16 or 24) should choose when I save my images in TIFF format? | A.The program gives you three options: 8-bit gray, 16-bit grey, and 24-bit RGB. What bit number you should use depends on what you want to do with your images. The camera is able to capture 12-bit intensity values. Therefore, saving your images as 8-bit gray values will scale your original 12-bit intensity values to fit 8 bits. If the only thing you intend to do with the images is to make figures using a graphics software package (e.g. Photoshop), then we suggest saving single channel images (only one fluorophore) in the 8-bit gray format. If you want to work with color images (e.g. when you have multiple fluorophores in the same image), then each channel (red, green, and blue) is limited to 8 bits and your only TIFF-format option will be the 24-bit RGB format. Finally, if you want to do intensity measurements in your images with analytical software such as NIH-Image, you may want to keep the true values registered by the camera. In this case the 16-bit gray format is preferable, but each wavelength in a multi-labelled specimen will need to be saved individually. |
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Q. How much space do DV users have on the server? |
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Q. What data files should I save, and what is the best way to save them? |
A.This is a judgement call and depends on how much space you have for saving images. Generally it's always good to save the raw data files (ON THE SERVER!!), since you can always deconvolve them again and make new TIFFs, etc. if you need to. Apart from this, just save whatever you're sure that you will use for your research and what you have space for. Realistically, biologists save everything. It's our nature. |
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| General Microscopy |
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Q. How can I study live cells? |
A.All microscopes in the CSIF are inverted, so cells must be in a leak-proof chamber with a coverslip/ coverglass bottom. Petri dishes are not optically clear and can only be used for low magnification (10x, 20x). We recommend the "LabTek II chambered coverglass" available from Nunc NOTE: Standard chamber slides which WILL NOT work as they have a glass slide bottom which is too thick to be imaged through. The chambers noted above have a coverslip/coverglass bottom (i.e. the thickness is approx. 0.17 microns). If you are using the Bioptechs heating stage, you can order the Delta T3 plates directly from Bioptechs http://www.bioptechs.com . CSIF has a limited amount of chambered coverglass and bioptechs plates for sale, so that you can get started on your experiments without waiting for your order to arrive. |
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Q. How can I mount a thick specimen for microscopy without sqashing it? |
A. What you need to do is to support the coverslip at a position at least as high as the level of your cells/sample. People have used a variety of techniques, including:
Use tape (scotch tape, electrician's tape) as "spacer" between slide and coverslip Make tape "well" by cutting out a small area in center of tape, put sample in well, add coverslip on top and seal. |
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Q. Can I fix GFP labelled specimens? |
A. GFP fluorescence is killed by dehydrating or fixation agents of ethanol and acetone, probably resulting from denaturing of the GFP molecule. Some groups have reported that GFP signal is also greatly reduced by formaldehyde fixation and that long term storage of samples in aldehydes will eventually wipe it the GFP signal and will increase autofluorescence. Other groups, however, have successfully fixed yeast cells with paraformaldhyde (4% in buffer, 1 hour fixation) without loss of signal. It is recommended, therefore, that you employ low percentage paraformaldehyde for the shortest periods of fixation as possible (5- 10 minutes for cells). Samples should be mounted in glycerol based mounting medium (approx 90% buffered glycerol).
Note: if your GFP is not a fusion protein that gets fixed in place, you will lose all signal upon permeabilization by detergents. There are reports that most nail polish (actually the solvent system of the nail polish, which is ethyl acetate/acetone) will lead to loss of GFP signal. Groups report success with the following: The best direct imaging of GFP is obtained in living tissue and cells. The addition of super radical scavengers in cell cultures systems is a good idea to help slow the bleach rate. |
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Q. What do antifade reagents do? |
A.When exposed to excitation light, all fluorescent dyes fade (photobleach). The photobleaching is dependent on: 1) the intensity of illumination; and 2) the duration of illumination. Other factors which influence the fluorescence intensity and bleaching of fluorophores include the pH and base solution of the embedding medium and the presence of other substances that quench fluorescence. The photon output of a dye represents the average number of cycles of excitation followed by fluorescence emission that the dye goes through before it is irreversibly photobleached. The average photon output is defined by the ratio of the probability that the dye will fluoresce (fluorescence quantum efficiency or Q f ) and the probability that it will photoreact irreversibly to become a nonfluorescent species (photobleaching quantum efficiency or Q b ). For example, fluorescein, which is very photolabile, has a Q f /Q b of about 30K in alkaline solution. Both Q f and Q b are properties of the dye that may be affected significantly by the dye’s environment. The primary environmental influence on Q b is the presence of singlet oxygen and free radical species. The main purpose of any antifade reagent is to sustain dye fluorescence, allowing longer observation times. This is usually accomplished by inhibiting the generation and diffusion of reac-tive oxygen species, thereby reducing Q b (preferably without any accompanying decrease in Q f so that fluorescence will persist). |
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Q. Which antifade reagents are recommended? | A.A list of a few commonly used (and most effective) antifade reagents and their advantages/disadvantages is listed below. If available, the commercial source is also provided. References and comments from Confocal Mailing List subscribers are also given.
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Q. What is the protocol for making DABCO anti-fade mounting medium? | A.DABCO embedding medium
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Q. What is the protocol for making N-propyl gallate anti-fade mounting medium? | A.5% N-propyl gallate in glycerol (w/v)
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Q. The signal collection is very weak. What can I do? | A.The single most critical parameter in determing the quality of the image is the quality of the specimen preparation. A good staining procedure should yield strong signal with an absolute black background. The following is a partial list of things to check if your staining is not optimal
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Q. Why is there spherical aberration if you use coverslips that are thinner than those recommended for the objective you are using (usually 0.17 mm)? |
A.Question on News Group: I never really understood that one, at least not for oil immersion lenses (Although I take care to get the right cover glasses anyway.) Shouldn't the immersion oil and the cover glass have the same refractive index so that the exact thickness of the cover glass shouldn't matter, at least in theory? (Water lenses would be a different story, of course.) Answer 1 (Lutz Schaefer): You still have to watch the coverglass thickness since your embedding media and sample likely have a refractive index that is smaller than 1.515 (glass, oil). Therefore most objectives produce the least spherical aberrations when the object lies directly on the coverglass. Take into consideration that for the condition that the sample refracts by 1.33 (Water) with the objective corrected in such way, a different coverslip thickness would shift the sample-to-coverslip distance that is necessary for aberration corrected imaging. In the older days with the 160mm tubus people could correct for that by changing the tubus length, but now with ICS we have to be a bit careful. Your point is that you assume that the RI of the coverglass is the same as that of the oil and that there is no interaction on the glass-oil interface. Unfortunatley very small differences in the RI's can become huge by large NA objectives (Answer 2). Contributing to that is also that depending on the working distance, the glass-oil surface is much closer to the front element of the lens than the glass-specimen interface. Very small refraction angles (or angle differences) at the glass-oil surface will cause a greater shift in the optical path than those on the bottom of the coverslip. For further information I suggest the following reference: "Experimental test of an analytical model of aberration in an oil-immersion objective lens used in three-dimensional light microscopy", S.F.Gibson, F.Lanni, JOSA Vol.9, No. 1/Jan. 1992 Answer 2 (Jim Pawley): Until recently, and as long as the specimen RI was about that of oil, I would have agreed. According to Ernst Keller's chapter in the Handbook, the oil (for Zeiss lenses) should have an RI of = 1.5180 ± 0.0004 @ 546 nm or 1.515 @ 589 nm (@ 20 deg Celsius) but the coverglass is 1.520 @ 589 nm. So the RI's are close, but not exact. Therefore, even oil objectives are only properly corrected for a certain distance into the specimen. (The error caused by incorrect coverslip thickness is, of course, even more severe for dry objectives and this is why so many of these have correction collars.) Furthermore, the change of RI with wavelength (dispersion) is different in the oil than in the Crown glass that the coverslips are (supposed to be?) made of. These all sound like small changes but, for high NA lenses, they are significant. Wilson and Juskaitis reported at the Focus on Microscopy meeting in Heidelberg last April that the process is so fussy that optimal lens performance was only attainable if the oil was within one degree C. of that for which its RI is specified (i.e., 20, +/- 1 deg C.) So watch out when you use oil lenses on specimens that must be at 37 deg C! On a modern microscope in which the tube length cannot be changed because of the design of the instrument, the only solution is either to purchase an optical element capable of changing (correcting?) the spherical correction for your system (e.g. an objective with a correction collar or a correcter from Infinity Photo-Optical (http://www.infinity-usa.com)) or to mix special immersion oils that will change the correction of a particular lens for specific circumstances. For instance, one might find that using oil with an RI = 1.520 permits reasonable correction when using an oil lens, and focusing about 15-20 microns into the larvae of a fly with a particular coverslip thickness and objective lens (or at least, a specific objective working distance, as this determines the actual thickness of the oil film used.) Of course, if the oil produces the "correction", this effect must be proportional to thickness of the oil and thus will become less as one focuses farther into the specimen (and moves the objective closer to the coverslip). At the same time, the aberration caused by the fact that cytoplasm has a different RI than oil (and than water!!) means that the needed correction becomes more just as the thickness of the oil is becoming less. As a result, the "special oil" solution works only over a limited focus range and this must be determined by trial and error for each specific case. Right now, it is my belief that spherical aberration is the main source of signal loss when viewing thick specimens with either oil or water lenses. This is probably because at least some cells seem to have an (average!!!) RI of somewhere near 1.45. This is far from the correct RI for either water or oil lenses, but fortunately the problem is only really severe when using high NA lenses (1.2 - 1.4) It is much reduced when using dipping lenses or 0.75 - 0.8, and not just because there is no coverslip to worry about. Mostly it is because it is the high NA rays that are most severely disrupted by the presence of a layer that is thicker or of different RI than that for which they were designed. |
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| Course Information | |||||||||||||||||||||||||||||||||||||
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Q. After taking a course, can I use the microscopes? | A.The courses are introductory one hour seminars and are intended only as an overview of the imaging technologies. In order to use the microscopes on your own samples, you must arrange for an individual training session. |
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Q. How do I sign up for a course? | A.To sign up for courses, put your name on the sign up sheets on the board outside Beckman B050 or e-mail microscopy@stanford.edu. Note that space is limited to 10 students per seminar, so if you e-mail, we may reply that the class is full. |
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Q. What is covered in "Introduction to confocal laser scanning microscopy"? | A.See description on Courses page |
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Q. What is covered in "Introduction to deconvolution"? |
A.See description on Courses page |
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Q. What is covered in "PhotoShop - from images to figures"? | A.See description on Courses page| Microscope manuals |
Here are links to downloadable manuals for the different microscopes that are available for independent use in the CSIF. |
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The manuals are in PDF format, so you need to have Acrobat reader installed on your computer. Note: These manuals cover the contents of the mandatory training sessions. If you have specific questions, please refer to the manufacturers' manuals that can be found by the respective microscope, to the frequently asked questions on this page, or ask the CSIF staff. | ||||||||||||||||||||||||||||||||
Almay Creme (hypo-allergenic, "Canyon" shade)
Vector Laboratories, Inc.