NIH Image : Use In Fluorescence and Confocal Microscopy

With many thanks to Wayne Rasband for his invaluable contributions to scientists of all disciplines, and his advancement of image processing.

The work that led to this manual was supported by grants to H. J. Karten from NIMH, NINDS and NEI. This manual was prepared with the direct support of NIMH and the Human Brain Project and the Brain Database Project. Under the stress of attempts to obtain grant support, we often neglect to sufficiently thank the many dedicated scientists who work so hard to establish those programs that provide the necessary funds. This manual is dedicated to those many loyal supporters of all of us at NIH, NSF, DOD, NASA, and elsewhere.

I have adopted and adapted a number of macros contributed by various NIH Image users. The authorship of the original macros was often difficult to determine. I am grateful to the unacknowledged authors, and hope that they are not offended if not specifically credited. My special thanks to Rusty Gage and Larry Goldstein for providing me with unlimited access to their confocal microscope facilities. Without their generosity, this manual would not have been possible.

Please let me know if this manual is helpful. Please inform me of any of errors in this manual. If you have any suggestions to improve this manual, additional macros that are helpful for processing confocal images, or other strategies for processing CLSM images, please let me know, and I will try to incorporate these changes in future versions.

Happy confocaling,
Harvey J. Karten, M.D.
hjkarten@ucsd.edu

2. Fluorescent Excitation/Emission: A Moving Target

Fluorescent excitation/emission is a degradative process. The brighter the excitation (within limits), the brighter the resulting emitted image. In order to examine the image for adequate evaluation of its quality and content, the user will expose the tissue for increasing lengths of time. Both excitation brightness and duration of exposure, while required for evaluation of the tissue, also degrade the quality of the fluorescence. At saturating levels, the half life of FITC and TRITC is only about 1-2 seconds. When dealing with in vivo or in vitro specimens, the problem is even more notable, as the prolonged exposure to intense excitation light degrades the cells, and hastens cell death. (This has been used to good benefit in the selective killing of identified cells in nervous tissue). Achieving good dark adaptation by the observer, use of efficient microscopes, supercooling the specimen, pulsing the light source and various reagents (often toxic to living cells) have all been used to diminish the fading, but the inevitable lose of fluorescence cannot be fully avoided.

As a result of these limitations, you may find that you are reluctant to examine the tissue at length before going to the confocal microscope to capture your "perfect" confocal images. This results in many bad confocal images, often consequent to the fact that the original tissue was not properly evaluated before using the CLSM. All the tricks in the world of post-acquisition processing of confocal images won't improve a lousy specimen. An expensive CLSM won't take a good picture of a bad specimen.

Far too often, investigators would be better served learning how to optimize their use of a fluorescent microscope more thoroughly before spending time hacking images from a confocal microscope.

3. Selection Of Fluorophores

This is a large and complex topic. It is dealt with extensively in the excellent chapter by Brelje, Wessendorf and Sorenson (1993) in the monograph Methods in Cell Biology: Cell Biological Applications of Confocal Microscopy , edited by Brian Matsumoto.

If working with a single fluorophore, most users seem to prefer either FITC or Cy3. FITC has long been used, and the emission wavelength corresponds to the range of peak sensitivity of the human eye. Cy3, with an emission wavelength in the short red range, has become increasingly popular. Cy3 is intensely fluorescent and bleeds both high and low--use it only in single fluorophore configuration, unless you have previously determined that the two compounds of interest definitely do not co-localize in the same structure. The great advantage of so bright a fluorophore is that you can use a smaller aperture, with a favorable signal to noise ratio, lower laser power and shorter exposure times.

My own preferences are:

A) Single fluorophore: Cy3

B) Double label fluorophores: FITC and LRSC or FITC and Cy5

C) Triple label fluorophores: FITC, LRSC and Cy5

D) Quadruple label: aminomethylcoumarin acetate (AMCA), FITC, LRSC and Cy5 (AMCA requires a laser that excites in the range of UV to short blue. While expensive, microscopes with lasers in this range are increasingly available.)

All these fluorophores can be obtained from Jackson Immunoresearch Laboratories, Incorporated (West Grove, PA).

4. Using NIH Image To Visualize Cy5

Many people have difficulty with Cy5 for the very reason that makes it so useful--it's emission is widely separated from that of FITC, and thus is barely visible to the naked eye. It is only visible if you are completely dark adapted, have a very efficient microscope and the intensity of fluorescence is robust. Even then, you will not be able to see much of the fine detail.

However, as users of NIH Image , you have one of the best tools available for imaging Cy5, if you have a video camera that supports on-chip integration (see following section) and a Scion LG-3 frame grabber. CCD video cameras are quite sensitive in the red to infra-red range. Wayne Rasband has provided a useful macro that allows you to vary the on-chip integration time and adjust the duration of integration while viewing the image on your computer monitor. For even greater control of signal fading, you should also use a Uniblitz shutter between the fluorescent light source and the microscope (see "Macros For Shutter Control" below).

5. On-Chip Integration

The intensity of fluorescent images is relatively low compared to the sensitivity of most CCD cameras. Video cameras sample available light, then send a complete image to the monitor every 1/30 of a second (33.3 msec or 30 frames per second). At that moment, the previous image is erased and the camera again starts to accumulate a charge proportional to the intensity of the incident light.

As the amount of light declines, the camera produces a more grainy, noisy image. With progressively lower light levels, the camera fails to detect a sufficient number of incident photons to provide a useful image. Averaging frames will not help, nor will software integration of multiple weak images.

In photography with film, the user has a number of options:

A) Increase the speed of the film

B) Increase the aperture or light collecting ability of the optics

C) Increase the exposure time--i.e., decrease the shutter speed

The analogous situation pertains to improving image capture with video cameras.

A) Increase the speed of the film

If you need to observe rapidly changing events, the only strategy you have available is to increase the sensitivity of your detector by using an intensified video camera. This is quite expensive, and the quality of the image is often degraded by the intensifier.

B) Increase the aperture or light collecting ability of the optics

There have been great improvements in the quality of fluorophores, light sources, lenses and filters. However, many images are still too faint to be adequately captured with a standard video camera.

C) Increase the exposure time--i.e., decrease the shutter speed

If your specimen is stable, then you can opt for longer exposure times.

On-chip integration acts by increasing the exposure time, thus increasing the number of photons captured. Rather than sending the image to the monitor every 33 msec, photons are allowed to accumulate on the sensor, and the image sent to the monitor when it is deemed adequately saturated. This technology has been available for many decades, but the cost often seemed beyond the budget of most labs.

NIH Image , in conjunction with some inexpensive hardware, now permits this technology to be widely used.

In order for this to work properly, you need four components:

A) A video camera capable of on-chip integration.

B) A source for a properly timed TTL pulse.

C) A frame grabber (video digitizer) that grabs the image at exactly the correct moment that the camera is sending the integrated image.

D) A computer and software that can control this process.

6. Practical Guidelines For Implementing On-Chip Integration

A) Choice of Camera: Several video cameras now provide built-in circuitry for on-chip integration at no additional cost. These include the two most widely used video cameras in labs, the Cohu 4915 (Cohu Inc., San Diego, CA) and the Dage-MTI 72 (Dage-MTI, Inc, Michigan City, IN). Images are allowed to accumulate on the camera sensor until a suitable level of exposure is achieved. A TTL pulse is then sent to the camera indicating that the computer now expects to receive the accumulated (integrated) image.

B) and C) TTL Pulse Source and Synchronized Frame Grabber: The Scion LG-3 card has made this technology available at low cost. Under normal operating conditions, the LG-3 continually samples the input from the video camera and displays the results to the monitor. However, under appropriate software control, the LG-3 card provides an output of a TTL pulse that is synchronized to the video digitizer, and captures the next image and holds it in the display buffer.

D) The software for controlling this whole process is provided in NIH Image . Wayne Rasband has provided a macro to control the duration of the exposure time for on-chip integration.

7. Macros For Shutter Control

I strongly recommend the use of Uniblitz shutter between the excitation light source and the specimen. The Uniblitz shutter is manufactured by Vincent Associates in Rochester, N.Y. Most microscope manufacturers can provide an adapter to mount the shutter between the excitation source and the microscope. The shutter is controlled by a Uniblitz controller box connected to the computer.

The advantage of using the shutter control is that it will limit the exposure of your tissue to the fading effects of the excitation by restricting exposure times to only that period required for actually collecting the images. Without this, I often forget to manually close the shutter, only to discover that I have burned out my best specimens.

Chi-Bin Chien, formerly of UCSD, has written a series of macros to control the shutter. I have combined the shutter control macros with Wayne's macro for on-chip integration. These macros are available as part of the collection of "Confocal Macros" provided at the NIH Image FTP site.

The following description pertains to the use of a fluorescent specimen. I suggest that you practice the use of on-chip integration with a transmitted light specimen, with the illumination set to a very low level. Once you have mastered the procedure for on-chip integration, you can then experiment with a fluorescent specimen.

A) A camera window should be selected. The shutter can be closed. Turn off the AGC (automatic gain control) on the CCU (camera control unit ). Set the manual gain and manual black level fully counterclockwise.

B) Select the macro for on-chip integration. If you do not have a Uniblitz shutter, you should open the manual shutter to allow excitation of your specimen.

C) If the image is too dim, move the mouse towards the top of the camera window.

D) If the image is too bright, move the mouse towards the bottom of the camera window.

E) The information window in the lower left corner of the screen will report on the number of frames that were integrated for the latest image. You can modify the macro to provide a direct readout of the duration of integration in seconds.

F) Observe the histogram for best spread of brightness values.

G) You may find it helpful to make minor adjustments in the quality of the image by changing the gain and black level settings on the CCU.

H) When you are satisfied with the quality of the image, terminate data collection by moving the mouse off the left side of the camera window and hold down the mouse button. The last image will continue to be displayed on the monitor. The shutter will then close, protecting the specimen from further fluorescent excitation.

This procedure requires practice to obtain images with a broad range of values.

As on-chip integration time increases, you will note a marked delay between the time you press down the mouse button and the appearance of a change in the image.

8. On-Chip Integration, Multiple Labeled Sections, And RGB Images

NIH Image permits you to evaluate the quality of multiply stained sections prior to using the confocal.

If you first obtain optimal quality images on the standard fluorescent scope using NIH Image , you can then combine the images to produce a useful RGB Stack. This can be tentatively evaluated using the NIH Image function RGB to Indexed Color . The Stack can also be saved and produces a high quality 24-bit RGB image that can be further modified in Adobe Photoshop.

A) Collect the image that you wish to be in the red slice of the Stack, using on-chip integration. Adjust the gain and black level to obtain the widest range of values, as indicated in a histogram of the image.

B) Run the macro provided "Make Stack from Current Image". This will take the current selected image, make a new Stack with three slices, for the R, G and B planes, and paste the selected image into the "red" plane.

C) Now collect a second image using the on-chip integration. Copy it to the buffer and paste it into the second slice ("green" plane).

D) If you have a triple labeled section, collect this with the on-chip integration routine, place it in the third slice ("blue" plane).

E) You can now use the RGB to Indexed Color operation under the Stacks menu to generate an indexed color image showing the double or triple labeled result.

F) Save the file as an RGB TIFF file. It can now be opened with Adobe Photoshop for better color images.

If you open a file on your a PC-formatted Zip disk on the Macintosh, and then Save it, you will have difficulty using the disk again on a PC. Iomega has not been helpful about the issue of cross compatibility of their drivers between PC and MacOS.

In view of the problems I have encountered with the Zip drives mentioned above, my current practice is to transfer the contents of the PC-formatted Zip drive to a magneto-optical with Macintosh formatting. If I want to use the files on a Macintosh (e.g., at home), I transfer the files to a Zip disk that has been formatted for the Macintosh.

The archival stability of Zip drives is still unknown. I strongly recommend that you store the original data files on archival media.

E) Magneto-Optical Disks For Archival Storage

The simplest means of storing large quantities of data is to store the file on a removable disk medium of large capacity that can be read by DOS, Macintosh and OS/2 systems. Since confocal images generate large data files, most systems have magneto-optical (MO) disks of 500 MB to 1.3 GB. A popular medium for this is the 1.2 GB MO disk, sold by Sony, Verbatim and others. These have the advantage of large capacity and have an archival life of at least thirty years. The most common drives are the Tahiti Max Optix 3, Pinnacle Sierra, various Sony, Ricoh, HP and NEC units. The disks are frequently (though not always) interchangeable. Many BioRAD confocal scopes were supplied with Panasonic MO drives. These use a proprietary disk that cannot be read by the drives of the previously mentioned manufacturers. When confronted with a Panasonic drive, use a LAN or FTP to transfer your files.

Until recently, these were the most common means of primary data storage for confocal microscopy. The price of the disks have dropped recently, and are now available for about $50 for a 1.3 GB disk. This is about half the price of a Jaz drive (also 1 GB), and though slower than the Jaz drives, is less prone to data loss.

The popularity of magneto-optical disks has declined recently due to the widespread availability and low price of Zip drives. Many people have had problems when attempting to read standard 1.2 GB disks that were originally formatted on a DOS based Pinnacle or Tahiti drive. This is particularly severe if you are using OS/2 with LaserSharp 1024. A significant problem with magneto-optical disks is the lack of universal driver standards. Use "Multi-Driver" on your Macintosh to facilitate reading DOS-formatted optical disks. "Multi-Driver" is a Macintosh Control Panel included in a software package sold by PC Access.

The BioRAD OS/2 optical driver cannot read Macintosh-formatted magneto-optical disks at this time. I no longer recommend using magneto-optical disks to move files from a PC platform (Windows or OS/2) to a Macintosh. Go with a Sneaker net using Zip drives, LAN or FTP. Magneto-optical disks, however, remain an ideal medium for archival storage of large files.

2. Opening BioRAD Files in NIH Image

BioRAD files consist of a 76-byte header that defines the size of the image in width and height, states if it is a Z-series, and, following the image data, provides information on parameters during collection of data, magnification scale and notes. The file structure of the new software, LaserSharp 1024, is similar to that of the earlier versions of Comos, with only minor exceptions pertaining to the footer at the end of the file that contains information about the parameters used during data collection.

Individual BioRAD files can be opened using the Import function of NIH Image . However, this requires that the user know the width and height of the individual file (Most frequently 768x512 in BioRAD files from an MRC600; 512x512 or 1024x1024 in LaserSharp 1024 for an MRC1000; 512x512 in Leica files). Unfortunately, I cannot figure out how to transfer a merged 8-bit BioRAD image to a Macintosh. The associated LUT (look-up table) is apparently stored in a manner or location that eludes me. If someone has solved this problem, please let me know. The Import function does not permit importing a Z-series. The macro for importing BioRAD files will correctly import a Z-series.

The macro "Import BioRAD MRC Z Series ", contained in the associated file "Confocal Macros", enables opening of either single sections or a Z-series. This works equally well on files generated with Comos and those generated with the recently released software, LaserSharp 1024. LaserSharp 1024 operates under OS/2. These files can be opened using the same macro employed to open the DOS-formatted files. LaserSharp 1024 stores each separate color plane of an RGB Z-series as a separate file. All three image planes of a single RGB file can be optionally stored as a single file.

After loading the BioRAD macro, you can "run" it by selecting it from the Special menu. The dialog will ask you for the "Starting Slice" of the Z-series. The default is 1.00. Accept that value for the moment. A BioRAD file containing only a single image will appear on the screen with the title of the original file (e.g., Axons03.PIC). I suggest that you immediately Save the file, prior to making any modifications, (e.g., "Axons03.PIC.Img"). This macro will load as many sections as possible, dependent upon memory.

If the number of Z-sections exceeds the memory allocated to NIH Image , the macro will stop running. You can Save the first group of sections in a separate file. Close the file, and again run the macro "Import BioRAD MRC Z Series", but when prompted for the "Starting Slice", enter the number of the last Z-section displayed. If the original Z-series is very large, you may have to do this several times. You can solve this problem either by buying lots of RAM, or store Z-series in smaller increments. This problem is particularly notable if you have collected images at 1024x1024 as well as collecting an extended Z-series. (e.g., 70 sections at 1024x1024 results in a file of ca. 74 MB.)

The associated calibration and notes of a single image file will be placed in a separate NIH Image text window. If you wish to include this information on the image that you saved as an NIH Image file, you must Paste it from the text window to the image window. If your notes contain information about pixel size and dimension of the image, use that to calibrate the scale of the image. (See "Set Scale" in NIH Image Manual ). Once you have calibrated the image, Save the file once again. The current version of the BioRAD macro does not properly read the calibration or notes file attached to a Z-series file.

BioRAD Z-series can be stored as a single large file, or as a series of individual sections. The former configuration has the advantage that all related files are stored in a single locus. However, it also means that such files may be huge. Using the above macro, a BioRAD Z-series will be opened into a new Stack. Save this in a similar manner suggested above for a single image file.

If this is a Z-series, use Stacks menu, and select Options to enter the step size of the motorized focus used when originally recording the Z-series. The magnification and slice spacing are stored with the original BioRAD file.

You may find it useful to Add a Slice to the beginning of a Stack to record specific comments about the series, what the data represents, and an image of Z-projection to demonstrate the contents of the file.

Save the original BioRAD *.PIC files, just in case. You can move all the original files into a separate sub directory to reduce clutter.

A) BioRAD Split Screen Images

Simultaneous collection of double labeled sections on the BioRAD can be displayed and saved on a "split screen" image, with the two images from PMT 1 and 2 displayed "side by side." This has two advantages:

1. You can directly compare the location of different antigens simultaneously
   

2. When collecting a Z-series with simultaneous split screen, you can be assured that the two images will be in the same focal plane.
   

The macro described in the previous section will open and display the split screen image. If you want to separate the two halves of the screen into separate images, use the macro "Merge BioRAD Split". This will convert the original split image into a three slice Stack (left image, right image and one blank black slice), then produce a merged color image. If you want to interchange the red and green planes, Select the window containing the Stack of three images, and run the macro "Swap Red_Green".

The Stack of three slices (left, right and blank), can then be saved as a 24-bit image in Adobe Photoshop 3.0 format, in the following manner:

1. If you have run the preceding macro, and have generated an RGB image in NIH Image , NIH Image will save the file as an RGB-TIFF file. This format is accepted as an Adobe Photoshop 3.0 RGB image.
   

2. Save this file with a modified title in order to preserve your original file.
   

3. Open the file using Adobe Photoshop 3.0.
   

B) Merging Split Screen Z-Series

The macro that operates on a Z-series of a split screen multi-labeled section will:

1. Place the left image in one Stack, and the right image in a second Stack.
   

2. Color Merge the two Stacks, and that will allow you to Animate the Stacks. This is very helpful when you want to track individual processes in a complex field through a lengthy Z-series.
   

   
   

Leica file format is a TIFF format. The Leica operating system is a peculiar hybrid using a VME bus with a 68040 CPU operating under OS/9, but uses a Windows front-end that generates DOS types of files. Leica has announced the release of a new software/operating system for their confocal microscope, based on Windows NT. The format of the images, however, will reportedly be unchanged. The original Leica files (and the new Windows NT files) can be directly read by a Macintosh. They will show up on your Macintosh desktop as PC files, with various optional icons, depending on how you set your system parameters in the Control Panel, PC Exchange. Each separate image file is accompanied by an "info.dat" file. Z-series are stored as single files with sequential numbers but with only a single info.dat file for the whole series. In order to reduce confusion in handling these Z-series with large numbers of files, move each Z-series set into a separate folder on your Macintosh.

If you are running NIH Image , you can Open these files directly, without having to Import them. However, you will not be able to double-click on the files to open them in NIH Image . In order to do that, pursue the following procedure:

A) Obtain a copy of "CTC 1.4" (or later) to change file type and creator. CTC 1.4 is available from the NIH Image server at zippy.nimh.nih.gov.

B) Select all the Leica graphics files in each folder (not the information.dat files) and drag them on top of the icon for CTC.

C) Using the resulting dialog box, change the "New Creator" to Imag , and the "New Type" to TIFF (creator and type are case sensitive, so enter exactly as spelled).

All the files will now have an NIH Image icon, and will be treated as NIH Image files by the program.

The info.dat files should be changed to "New Creator" Imag and "New Type" to TEXT . This file contains important information including scaling factors, size of image, size of pixel, and spacing in a Z-series.

To open a Leica Z-series Stack:

A) Close all other NIH Image windows at this time. Make sure that you have placed all the related Leica image files into a separate folder.

B) Select Open from the File menu. In the resulting dialog box, go to the desired folder and click "Open All", then Open . This will open all the files in rapid sequence.

C) Select Windows to Stack from the Stacks menu. Make sure that you set the magnification/calibration scales and slice spacing.

D) Save the resulting Stack with a new name.

4. Zeiss Files

Zeiss files saved using the newer version of software with the LSM 310 and 410 are "standard" *.TIF files. Use the same procedure outlined above for the Leica files--i.e., use CTC to change "New Creator" to Imag and "New Type" to TIFF . The files can then be directly opened by NIH Image . Make sure the LUT is correctly set. I have often found the Zeiss LUT map to be inverted, with a negative slope, and an inverted image. Click on the icon in the lower left corner of the mapping window to obtain the correct black/white relationship.

5. Molecular Dynamics/Sarastro

Jay Hirsh kindly provided information on the format of Molecular Dynamics files. The data files are simple files without a header or footer. Each image of a Z-series is stored as a separate file. Information about a single image, or about a Z-series is stored in a separate text file. The text file can be directly opened by NIH Image .

A) Place all associated images of a single Z-series in a single folder. Make sure that they are correctly numbered to reflect their order of collection (e.g., 008.ext, 009.ext, 010.ext, not 8, 9, 10).

B) Select Import from Stacks menu. The user must know the image format (512x512 or 1024x1024) in order to use this correctly. The offset value is 0.

C) Check boxes "Open All" and "Invert".

D) Click Open . This will open all the files to the screen.

E) Select Windows to Stack from the Stack menu.

F) Save the Stack with a name of your choosing.

6. Noran Confocal Files

Learn to use the adjustments to black level and gain on the CLSM to optimize the spread of gray values. If you have to trade off between using the "+1 to +3 LUT" position on the BioRAD versus a brighter setting of the laser, choose the brighter laser setting. It will burn out your specimen faster, but give you a better signal to noise ratio and a wider dynamic range in gray scale value. The "Photon Counting" mode, or the "Accumulate" mode on the BioRAD will often provide an image with the best dynamic range. On the BioRAD, I prefer to use the "Accumulate" mode with "Slow Scan" for best results. It is not the purpose of this manual to teach the use of CLSMs, but I only wish to emphasize the importance of the quality of the original image.

B) Avoiding High Contrast Images

If you get into the habit of checking the histogram of your images as you collect them on the CLSM, you will improve the quality of the original images, and find less need to twiddle with the LUT values.

C) Avoiding Noisy Images

If possible, collect images using the "F1" (Slow) setting (on the BioRAD) with a Kalman setting of at least 3, or "Accumulate" mode. When using "Accumulate mode", you can use a less intense laser source, and manually accumulate until the image appears satisfactory.

2. Editing Image

A) Cropping Images, Erasing, Superimposing Text, Scale Bars, Rotating And Scaling Images

NIH Image provides a wide range of tools for editing your image. These are described in NIH Image manual About NIH Image .

1. Notes On Scaling
   

1. Caution On Scale And Rotation Of Images
   

3. Using LUTs

The following operations are commonly used to enhance the image. These are standard operations on all confocal scopes, and are based on methods that are widely used for manipulating digital images. See appropriate section of the NIH Image manual.

A) Modifying Brightness And Contrast

In the simplest operation involving look-up tables, you may choose to emphasize a selected range of index values (gray scale values), and minimize other values. The choice may be based on your interpretation of the information content of the image. Learning how to interpret the histogram, and modifying brightness and contrast are essential skills in image processing.

B) Linear And Non-Linear LUTs, Including Custom LUTs

The standard LUT shown in the mapping window (lower left side of your screen) is a linear LUT. You can modify the brightness and contrast values by either dragging the Brightness and Contrast slider buttons, or by directly dragging the plotted line in the mapping window.

The "Confocal Macros" file included with this manual provides a number of alternate, non-linear LUTs based on sampling the values within your display buffer. These include Log, Parabolic, Square, Square Root and Gamma transforms. Play with each of these and observe the effects on your images. You will probably find the Gamma transforms most useful when used with a setting of 1.5 to 2.0. Many of the resultant changes produce results similar to those obtained on the BioRAD with +1, +2 and +3 LUT settings. When you open BioRAD files with NIH Image , the results may not match what you saw on the BioRAD. Most commonly, the image will appear much darker, and lacking the detail you so clearly remember seeing on the screen when collecting the original image. NIH Image has not corrupted your files. This is most frequently due to the fact that the image you viewed on the confocal microscope may have had a non-linear output LUT attached to it. The original data is imported intact, but the output LUT is not attached to the new NIH Image file, and a new linear LUT is appended to the file. See the NIH Image manual for instruction on how to modify the LUT.

C) Enhance Contrast Operator In NIH Image

The Enhance menu item in NIH Image has a specific function entitled Enhance Contrast. This produces a custom linear LUT effect that is as good as any result I can obtain in my attempts to manually modify the LUT curves. However, you may find that this operation produces an excessively contrasty image, with too steep a slope in the LUT mapping window. If so, then manually change the LUT in the mapping window to give a slope of the LUT about halfway between the original value and that produced by Enhance Contrast . If that is satisfactory, then Apply LUT (Command+L ). If you feel that you want still more contrast in the image, repeat the above sequence.

In order to appreciate the effects of this operation on the original image, examine a histogram of the image before and after this procedure.

As in all alterations to the LUT, the Enhance Contrast operation only modifies the display buffer, not the image in the file buffer or the original file on the disk. In order to alter the file buffer, you must perform Apply LUT . This will not alter your original disk file. If you wish to do so, Save the file. Once you have done that, you cannot go back to the previous image. You may, therefore, prefer to save the modified file under an alternate name.

Enhance Contrast is very different from the Equalize operation, which is likely to result in excessively splotchy images. Compare the effect of each of these operations on the appearance of the mapping, histogram windows and LUT.

D) Thresholding And Density Slicing

See NIH Image manual.

E) Pseudocoloring Images

Pseudocoloring confocal images assists the viewer in displaying double labeled sections, detecting major differences in concentration, or changing concentrations, as in calcium ratio imaging. See appropriate menu item, and NIH Image manual for use of this operation. All the color applications described in this manual, although they may resemble the original colored fluorescent image, are pseudocolor.

F) Exporting To Adobe Photoshop

Recent versions of NIH Image save single files and RGB files in a format that can be directly opened by Adobe Photoshop 3.0. This will not work with Z-series.

4. Enhancing/Filtering Image

NIH Image provides a wide range of tools for enhancing and filtering images, including filters to sharpen, smooth, shadow and detect edges. You can write your own kernels, run median filters, and Sobel operators. Some of these operations are multistage operations and alter both the display buffer and the file buffer (but not the disk file), and cannot be Undone . You will have to reopen the original file to restore the image.

For further details, see NIH Image manual.

5. Quantitative Measurements

NIH Image was originally developed for quantitative measurements. There are many useful functions described in the About NIH Image manual. Various measuring functions include: length, area, density, particle counting, and profile of a Line. About NIH Image will provide guidance in the use of these operations.

6. NIH Image Macro Language

Two color plate: If you wish to combine only two plates, open the two files.

Use the macro "Color Merge Two Images" contained in the sample "Confocal Macros" that is provided with this manual. If you examine the sequence of commands in the macro, you will find the following operations:

A) Open a fresh stack

B) Paste the "red" image into the first slice

C) Add Slice to add an additional slice

D) Paste the "green" image into the second slice

E) Add Slice to add a black (empty) third slice

F) Merge from RGB to 8-bit color using a custom LUT

G) Open a new window containing the merged color image

Alternately, you can do this operation manually to familiarize yourself with the procedure. The first file opened will become the red plane, the second file the green plane. Under menu item Stack choose Windows to Stack . This will put the two plates into a stack labeled "RGB". Any further operations will not alter your original data files, so you can always go back and start again.

Save a copy of the NIH Image Stack. The following section describes how to use this Stack with Adobe Photoshop 3.0.

If you want to alter the contrast or brightness of either of the color planes, you have to modify either the original images or those in the RGB stack. I suggest that you limit your initial attempts to the slices in the RGB stack. When you change the Brightness or Contrast of any single image, you must then Apply LUT (Enhance menu) to that single image. Do not use the Apply LUT to "Stack" macro as this will modify all the images in the Stack.

From the Stacks menu select RGB to 8-bit Color. In the resulting dialog box, select "Custom Colors". This will generate an "Indexed Color" window from the Stack, but will not alter the stack itself.

Since each original file may have a different distribution of values (see histogram), the saturation of each color plane may differ markedly. Be creative, try different ways of doing it. (e.g., use Enhance Contrast operation on one slice at a time). Try non-linear LUTs, various filters, etc.

The resulting "Indexed Color" image can be saved with a unique name.

Once you have mastered this sequence, you will have a clearer understanding of the operation of the macro "Color Merge Two Images" contained in the sample "Confocal Macros" that is provided with this manual.

If you now Save the RGB stack (RGB TIFF) in NIH Image , the resulting file can be viewed with Adobe Photoshop as a 24-bit file.

1. Merging Files With Adobe Photoshop
   

Version 1.58/1.59 of NIH Image now permits Stacks to be directly saved in Adobe Photoshop 3.0 TIFF format.

A) The stack must consist of 3 slices.

B) Before saving the stack, open Slice Info in the Stacks menu. Confirm that RGB is selected.

C) Save the file. It will write the file as an RGB TIFF file.

You can then directly open this NIH Image Stack in Adobe Photoshop 3.0.

An alternate means of converting the stack to an RGB TIFF format is to call the RGB to Color function. This will automatically change the "Slice Info" window to RGB. The resulting "Indexed Color" image will give an approximate (8-bit) preview of the results to be obtained with Adobe Photoshop (24-bit). Save this file.

You will now be able to use the various Adobe Photoshop tools to modify the resulting 24-bit image. You should explore the use of the Levels command (Command+L ), as well as the Brightness/Contrast command (Command+B ).

If you have a quadruple (or more) labeled section (e.g., 512x512, four fluorophores, or three fluorophores and a Nomarski/DIC image), and have stored them in a four slice stack in NIH Image , you can open them in Adobe Photoshop using the method described below.

A) From the Adobe Photoshop File menu, select Open.

B) Using the Adobe Photoshop Open dialog, go to the desired directory containing the NIH Image Stack file of interest.

C) Using the same dialog box, pull down list at the bottom of dialog box, choose Raw file format.

D) This will show all files in the chosen folder.

E) Select the NIH Image Stack containing the four slices. You have to know the dimensions of the slices.

F) The resulting dialog box requests that you fill in:

1. Width (e.g., 512 pixels)
   

2. Height (e.g., 512 pixels)
   

3. Number of channels (e.g., 4, for the number of slices in the stack)
   

4. Select "not interleaved"
   

5. Offset =768 (3 X 256 = 768)
   

H) Save the new Photoshop image using the Save As... dialog box. Do not save as a Raw image. Select TIFF format, and choose a new name for the file, otherwise the simple Save command will overwrite the original NIH Image file and store the data in the Raw format.

You can modify each color channel separately using several different methods, as described in the Adobe Photoshop 3.0 manual.

The quality of the resulting color image will generally be much better than that provided by NIH Image , as it can utilize a full 24-bit look-up table, and does not require dithering of the image.

The Adobe Photoshop manual and tutorial will provide further guidance in modifying the images.

1. Preferred Method
   

C) Compare Results Of NIH Image 8-Bit Merge With Photoshop 24-Bit Merge

The quality of the color image (assuming your monitor is set for 24-bit color) is generally much better in Adobe Photoshop than the optimized 8-bit image obtained in NIH Image .

D) Adjusting Color Contrast On Sections In A Stack (See Macros)

Do the Enhance Contrast operation separately on each section in the stack. Failure to do so will result in the merging of both saturated and unsaturated images.

If you are dealing with a single RGB section, you will find that Adobe Photoshop provides much better tools for this.

E) Merging A Double Labeled Pair Of Z-Series Using A Macro

1. Open the two Stacks. The first stack will be assumed to be the "red" stack. The second stack will be the "green" stack.
   

2. Close all other windows.
   

3. Select the macro "Color Merge Two Stacks".
   

4. Lean back and watch the fun.
   

The drawback of this procedure is that the resulting images are 8-bits, not 24-bits. I cannot find a way to do this procedure with Adobe Photoshop.

2. Projection Of A Z-Series And 3D Rotations

Many of the most valuable qualities of confocal Z-series are that the data can be manipulated to obtain "through views" of the stack, the stack can be resliced at various angles converting a transverse view into a sagittal or horizontal plane, it can be used to generate 3D images, stereo pairs, and other operations. Indeed, NIH Image is able to perform operations as well as much of the software provided by the manufacturers of confocal microscopes, and equivalent to many of the basic operations provided by very expensive image processing programs such as VoxelView, VolVis and VoxBlast running on a Silicon Graphics Workstation. For more elaborate operations using true Voxels, complex shading and ray-tracing in 24-bits, these latter programs are excellent choices. For the most common operations, however, NIH Image is quite adequate. Optimal performance on these tasks will be achieved using a PowerPC with sufficient memory to handle your typical data sets.

A) Stepping Through A Z-Series Using Stacks

1. Open a Z-series in a Stack.
   

B) Animating A Z-Series

1. From the Special menu, select Video for oscillating movies. This will produce smooth back-and-forth motion of the Z-series animation.
   

2. Command to Animate a series. Number keys 1-9 control the speed of the animation.
   

This is an extremely useful benefit of the confocal microscope. A stack of Z-series sections are all in optimal focus. If projected onto a single plane, all objects throughout the thickness of the imaged section will now be in sharp focus and spatial relationships may be more evident.

NIH Image version 1.58 and later provides macros to perform the Z-projection. A sample macro to accomplish this operation is included in the accompanying macro files.

1. Selection Of Area For Z-Projection
   

   A) Choose an extended Z-series (e.g., more than 10-20 images) with well defined profiles of objects.
   

   B) Run the macro "Project Z Series".
   

C) Use Project command

D) Set :
Slice Spacing (Pixels): Set with your slice spacing
Initial Angle (0-359): 0
Total Rotation (0-360): 0
Rotation Angle Increment: 0
Lower Transparency Bound: 0
Upper Transparency Bound: 100
Surface Opacity (0-100): 0
Surface Depth-Cueing (0-100): 100
Interior Depth-Cueing (0-100): 0

E) Select "Minimize Window Size"

F) Select "X-Axis"

G) Select "Brightest Point"

After you develop a sense of familiarity with the program, play with different values. Start with the default values. Once you are more familiar with the Thresholding tool (Magic Wand), you will be able to use that to set the range of values desired for your Z-series.

This will generate a single Z-projection of all the plates in the Stack, viewed from directly in front of the stack.

Experiment with different view angles by using different values for the "Initial Angle", while leaving "Total Rotation" at 0.

If you want to make a quick and dirty stereo pair of images from a Z-series, then the values should be something such as:

Initial Angle (0-359): 356
Total Rotation (0-360): 8
Rotation Angle Increment: 8

Select the "Y-Axis" as your "Axis of Rotation". This will cause the resulting image to rotate left to right (i.e., around the Y-axis). Choose "Brightest Point" as the "Projection Method".

This will produce a new Stack containing two images at an 8 increment. This is a common angle for stereo pairs. In order to visualize this, you can do two different procedures.

1. From the Stacks menu, Animate the stack, and the image will rock back and forth around the Y-axis.
   

2. Make a Montage using this function from the Stacks menu. You can experiment with different values for the scaling factor. The smaller the scaling factor, the easier you will find it to fuse the images. Ideally, for people with limited experience in "Free Fusing" stereo pairs of images on the screen, matching points on the screen should be less than 50 mm apart. With further experience, you will be able to fuse larger images with matching points further apart.
   

3. Optimizing Settings For Project Function Of Z-Series
   

   A) Make sure that "Invert Pixel Values" are not selected in the Edit/Preferences menu item. If selected, de-select this item. Then Select File/Record Preferences. Quit NIH Image in order to implement the changes in preferences. If this item is "selected", you will be easily confused by the various values displayed in the info, mapping and histogram windows.
   

B) There are a number of parameters in the Project item under the Stacks menu. Understanding the proper use of this item is critical to obtaining pleasing results in Z-projections and rotations.

C) The first of several items include obvious settings regarding "Slice Spacing", "Initial Angle", "Total Rotation" and "Rotation Angle Increment". These values are all obvious and pertain to the sampling interval of the resulting projection.

The remaining items on the list, "Lower" and "Upper Transparency Bounds", "Surface Opacity", "Surface-Depth Cueing", and "Interior-Depth Cueing", however, are very confusing, and cannot be easily understood without some explanation.

One of the greatest difficulties in mastering these functions is because once you have selected specific settings, obtained an image, and then try other settings, your screen will be cluttered with images, and the particular parameters used to obtain them has been forgotten.

Start with the "Lower" and "Upper Transparency Bounds" in relationship to the stereo pairs you generated a few minutes ago.

When making a simple single view Z-projection, the slice spacing is irrelevant. However, depending upon the setting of the "Lower" and "Upper Transparency Bounds", and "Interior Depth-Cueing", or the resulting image is likely to appear somewhat lacking in brightness.

"Lower Transparency Bound" must be set to 0. (Note that this value is expressed from 0-254). This value determines the cut-off point of displayed pixels. All pixel values below the setting selected will be discarded and modified to a darker value. Remember the whitest/brightest pixel on the Macintosh has a value of 0, the black pixel a value of 255. "Lower Transparency Bound" greater than 0 (e.g., n ), will modify all pixels from zero to n in your image and replace them with darker pixels. This results in dark gray to black holes in the midst of a bright object. The setting of this value is quite simple. The most effective way to appreciate the consequences of this choice is to look at histograms of the images produced with "Lower Transparency Bound" set to zero, and compare that with a histogram generated with a "Lower Transparency Bound" of perhaps 50. Make sure that your starting image has prominent objects with histogram brightness index values of 1-10 (out of a range of 0-255).

For the "Upper Transparency Bound", I suggest that you start with a value of 254. After using this for a while, play with other values.

The three following functions in the Project dialog box are expressed as values of 0-100%.
Surface Opacity (0-100): 0
Surface Depth-Cueing (0-100): 100
Interior Depth-Cueing (0-100): 0

NIH Image 1.61 sets "Interior Depth-Cueing" to a default of 50. This will produce "muddy" images. You must set this to 0.

D) Reslicing The Z-Series Along Alternate Planes: (X-Z, Y-Z And Theta-Z)

One of the most useful features of NIH Image is the ability to rapidly reslice a Z-series data stack along alternate planes. In addition to simple orthogonal planes (i.e., X-Z and Y-Z), you can reslice at any arbitrary angle between X and Y to generate a single Z-plane cross section. This is very useful for evaluating the extent of penetration of antibodies into tissue, evaluating cell spacing, etc.

The closer the images in the original Z-series, and the greater their number, the more natural appearing the final results. In order to provide a seemingly continuous image in the resliced plane, the software interpolates gray values. For improved quality of resliced images, your original Z-series should be as close as possible (e.g., 0.25 m).

Have you set the calibration for magnification and slice spacing, as described above? (Also see NIH Image manual).

Your original data file may contain the needed information about slice spacing. In many of our files, the step size was 0.38 m (on a BioRAD), and the pixel size 0.105 m (9.5 pixels per m). However, this depends upon the Z-axis step size, the objective and the zoom factor.

A) From the Stacks menu, select Options . Enter the appropriate value in the slice spacing dialog.

B) Select an image from the series that best shows the features of interest. Using the Calibration/Measurement tool (dashed line in the right hand column of Tool Palette), draw a line across the section along the desired plane of resectioning. If you wish to constrain the plane to either the X-or Y-axis, press the Shift key and hold down as you draw the line.

C) From the Stacks menu, choose Reslice (Command / ). If circular profiles appear excessively flattened in either the X- or Y-axis, experiment with different values of "Slice Spacing" in the Options dialog.

E) Reslicing To Make A Z-Series In An Alternate Plane

The preceding instructions describe how to obtain a single reslice in the X-Z or Y-Z plane. If you would like to obtain a new stack of Z-series section, not just a single section, use the macro provided, "Reslice Horizontally" or "Reslice Vertically". The reslicing is limited to orthogonal planes of section. Thus, if you wish to reslice along an oblique angle, you will first have to rotate the stack. In order to do this, use the Angle Measurement tool from the Tool Palette. Measure the degrees of variance of the object in the image from the angle you finally desire.

A) Use the macro "Scale and Rotate Stack", using the angle value established in the previous paragraph.

B) Now select a rectangular area that encloses the region you want to reslice in either horizontal or vertical plane.

C) Use the macro "Reslice Horizontally" or "Reslice Vertically".

1. Rapid Dynamic 3D Reslicing

Norbert Vischer of the Netherlands has recently contributed an extremely useful macro, "3D Slicer" for reslicing a stack of sections along the X-, Y- and Z-axes. This macro is provided in the program "Object-Image 1.59", available from the NIH Image FTP site. Object-Image 1.59 provides extremely rapid reslicing in real time. The individual resliced sections cannot be saved, for unlike the macros described above, they are not placed in a new stack. The reslicing of sections is accomplished by dragging the mouse along the X-, Y- or Z-axes of a master stack, and results in rapid generation of resliced images in the two other planes. Reslicing can only be done along orthogonal planes.

A) Open a Z-stack of sections, such as the sample MRI Images of a human skull and brain.

B) Enter the correct magnification scale and slice spacing (5.0 mm), as described above.

C) Under Stacks menu, select 3D Slicer. This will open a new window containing an angled perspective view of the original stack, with 3D-projection planes along the two alternate planes, as well. In the upper part of the window will be an image of the resliced section parallel to the X-axis. On the right side of the window will be the resliced section parallel to the Y-axis.

D) Place the mouse over the X, Y or Z margin. The cursor changes to a two headed arrow indicating the direction of movement. Drag the X-, Y- or Z- axis on the main central image, and observe the rapid resectioning in the alternate planes.

E) Place the mouse over the intersection of the X- and Y-axes and the cursor becomes a four-pointed arrow. You can now simultaneously reslice parallel to both the X- and Y-axes.

F) Double clicking the mouse on the X-or Y-axes will turn off the reslicing tool for that axis. Double clicking on the dashed lines on the X- or Y-axis will restore the reslicing planes.

F) Generating A 3D-Series From A Z-Series

This is a computationally intensive procedure. A fast PowerPC will prove very desirable when doing this operation. When you are still learning the basics of this procedure, use the Selection tool from the Tool Palette and outline a small portion of the image when generating a 3D-series. It will complete the task much more rapidly.

Use care in settings. Remember to correct for thickness and spacing of individual sections.

1. Select Project from the Stacks menu.
   

1. Set:
   Initial Angle (0-359): 0
   Total Rotation (0-360): 15
   Rotation Angle Increment: 180
   

Select the "Y-Axis" as your "Axis of Rotation". This will cause the resulting image to rotate left to right (i.e., around the Y-axis). Choose "Brightest Point" as the "Projection Method".

Now click OK and wait. The program will generate a new stack of images of the "rotating" objects in the original stack.

Once you have had further experience with this procedure, Project using a full 360 degree rotation at closer intervals, changing transparency values, axes of rotation and the various other options available.

A macro to facilitate this operation is provided.

G) Animating A 3D-Series (Producing "Apparent Rotation")

Under Special menu, select Video Options . Check "Oscillate Movies". Close the dialog box.

To generate the impression of rotation with your newly generated 3D stack, use the Stacks menu and select Animate (Command+= ). This is heavily memory dependent, so keep your initial efforts small.

You can control the speed of rotation with keys 1-9. You can step through single sections using the < and the > keys.

H) Make A Stereo Pair Or Series Of Stereos?

There are several methods of generating stereo images from a stack of sections collected in the Z plane. One simple method, the "pixel shift" method, is that used in the original BioRAD software.

The pixel shift method starts with a stack of a Z-series. You will generate two Z-projections from this original stack. One will be "pixel shifted" to the left and a second "pixel shifted" to the right. This is achieved by shifting each successive section in the stack by one additional pixel (or more, if desired) prior to performing the Z-projection. The two resultant images are then placed side by side. The pixel shift method is generally limited to stereo images centered around the original plane of image collection.

Projecting and ray-tracing is a second method that projects (ray-tracing) the stack onto an imaginary view plane from different angles of view and generating a 3D rotation series. This second method is computationally more complex, but provides the possibility of generating stereo views from any angle around a central point.

Start with the original Z-series Stack. Select the Project from the Stacks menu and set "Initial Angle" to 354, "Increment Angle" to 6 and "Total Rotation" to 12. This will produce a new Stack containing three slices (354 , 0 and 6 ).

1. Stereo Series In Black And White

Once you have generated a 3D-series, make a Montage series using the appropriate function from the Stacks menu. The Montage function is non-destructive, i.e., it does not erase the 3D stack, however, I suggest that you Save your work as you go along.

A) Stacks Menu: Montage

The resulting dialog box will show a series of values that are dependent upon the number of sections in the stack. If you made a simple stack with only 3 sections, then choose 1 row and 3 columns, and an increment of 1. If you generated a full 360 rotation series, you can select any range of slices (e.g., slices #4-8 out of 16 slices), or incremental slices (every 2nd or 3rd slice in the stack), and define the numbers of rows and columns you wish to see displayed.

This will generate a series of images in a new window on the screen. If the resulting images are too large to line up next to each other, use a scaling factor when generating the montage.

Start with two side-by-side images. To facilitate seeing the stereo effect without special viewers, choose a set of images with a prominent object to provide an alignment cue in the center of each image. Set the image size and separation with the alignment cues no more than 50 mm apart on the screen. Gradually work your way up to greater separations and then slightly larger images. Initially, you may find that you require a stereo viewer to visualize a stereo image from side-by-side images. With practice, you will not require a stereo viewer and should be able to scan across pairs of sections and jump from one stereo image to the next. (The typical interpupillary distance of most people is about 65 mm. You can also practice learning to fuse stereo images by using some of the recent popular books with "random dot stereograms", such as The Magic Eye .)

2. Stereo Pair Of Single Labeled Section In Color

An alternate manner of presenting stereo images is to merge a stereo pair into a single image plane, assigning one image to red and the other to green.

To generate a 3D rotation series around the Y-axes., limit the number of image to two at a separation of ca. 6-12 . For the first trials, set the "Initial Angle" to 354, the "Increment Angle" to 6 and the "Total Rotation" to 12. This will generate three plates in a new stack. Animate the new stack of images, and if the animation produces the desired effect, delete either the middle or one of the end plates. Add a black plate as slice 3/3. (See section above on Merging Slices to generate 8-bit color images). Using the Stacks menu item RGB to 8-bit Color , you will obtain a single merged image which can be viewed with a pair of Red/Green stereo glasses.

The greater the separation between plates (i.e., 12 rather than 6 ) the more exaggerated the stereo effect. Play with different values. Some people find that angles greater than 15-18 are excessive.

3. Color Stereo Images Of A Double Labeled Section

Although this is a more complex set of operations, it is a logical extension of the methods described above.

First generate a 3D rotation series of each Z-series of a double labeled section. Save the results. Close all windows except those of the two 3D rotation series. Now use the macro "Color Merge of Two Stacks". You will now have a rotation series (3D) of two simultaneous different fluorophores. Animate the series.

Now generate stereo pairs, using the montage method described above.

I) Exporting Stacks to QuickTime Movies and VCR Recordings of Stacks

A rotating Z-series, or any other Stack, can be saved in QuickTime format, allowing it to be viewed with a variety of other programs, such as a simple QuickTime Viewer, or with a program designed for preparing and transcribing clips on a VCR, such as Avid VideoShop 3.0. The latter program is presently distributed at no additional cost with many Apple Computers. Avid Videoshop can be used to prepare a VCR tape of your data, merging different data sets into a single sequence, etc.

1. Saving Stacks In QuickTime Format
   

   A) Save the Stack in standard NIH Image format.
   

   B) Open the Stack, and select Save As...
   

   C) There are a series of optional buttons at the bottom of the dialog box, including TIFF, PICT, PICS and others.
   

   D) Choose "PICS"
   

   E) Rename file with new name (e.g., if file name was "NewCell", rename it "NewCell.PICS") to avoid overwriting your original data file.
   

   F) The new file can still be opened as a Stack within NIH Image .
   

   G) If you Save the file, it will revert to the original TIFF format, though with the new filename, and is likely to be confusing when you try to open it with a QuickTime player. Therefore, any additional Save operations should always be done using Save As... .
   

1. Transferring NIH Image RGB images to Adobe Photoshop 3.0
   

2. New macro to automate Z-projection
   

1. Use of NIH Image for pre-evaluation of fluorescence--shutter and on-chip integration control. Provides a series of macros for controlling the shutter, on-chip integration, making new Stacks and related operations.
   

2. Database storage of images--put Z-projection image on top of Stack for databasing. Compare FileMaker Pro, Search 3.1, Cumulus 2.5, and Kudo.
   

3. GetPath functions--a means of imprinting the file name and full path on each image is provided in NIH Image version 1.60. This greatly simplifies record keeping.
   

4. Notes on Z-projections
   

5. Caution on rotation
   

6. Revision on recommendation for storage media.
   

These macros are arranged in order of my personal preference (more or less). This order will probably not satisfy anyone else. If you rearrange the order of these macros, make sure that you are careful in regard to defining "Variables and Procedures". They must all be placed in the file in front of the macros that call them.

Define Procedures for Stacks
procedure Error(s:string);
procedure CalibrateImage
procedure CheckForStack;
procedure CheckForSelection;
procedure GetSliceRange(smart:boolean);
procedure GetSliceSequence(smart:boolean);

Some Universal Operations
macro '[P] Print Video';
macro '[Q] Get Path';
macro '[F2] Calibrate Image';

Operations on Confocal Files: Import, Split, Merge
macro [=] "Animate";
procedure ShowBioRadInfo(InfoOffset: integer);
macro '[F1] Import Biorad Z Series';
macro '[S] Merge Split BioRAD';
macro ' Color Merge Two Images';
macro ' Color Merge Two Stacks';
macro 'Separate SplitScreen Z Stack';
macro '[F5] RGB to Indexed ';
macro '[W] Swap Red_Green';
macro ' Merge Two Stacks';

General Stack Operations
macro ' Make stack size front image';
macro '[F8] Flash to Stack
macro MakeStack w_Current Image';
macro 'Add Slice';
macro '[A] Add Black Slice';
macro '[D] Delete Slice';

Shutter Controls and On-Chip Integration
macro ' Autoshutter';
procedure ResetShutter;
macro '[O] Open Shutter';
macro '[C] Close Shutter';
macro '[T] Trigger Shutter';
macro '[R] Reset Shutter';
macro '[G] Open Shutter-Grab-Close';
procedure EndIntegration;
procedure Integrate (mode:string);
macro '[F6] Integrate On-chip Using Cohu';
macro '[F7] Integrate One Image on Cohu';
macro '[2] SetIntegrate:2 Frames';
macro '[F] Show nFrames';
macro 'Integrate White Light';
macro ' Live';
macro ' Average';
macro '[F1] Import Biorad Z Series';
macro '[F2] Calibrate Image';

Stack Modifications, Z-Projections, Stereo Pairs
macro ' Smooth Stack';
macro ' Sharpen Stack';
macro '[I] Invert Stack';
macro ' Reduce Noise Stack';
macro '[L] Apply LUT to Stack ';
macro '[F0] Remove 0 and 255 from Stack';
macro ' Flip Stack Vertical';
macro ' Flip Stack Horizontal';
macro ' Clear Outside Stack';
procedure CropAndScale(fast:boolean; angle:real);
macro '[E] Crop and Scale-Fast ';
macro ' Crop and Scale-Smooth ';
macro ' Rotate Left';
macro ' Rotate Right';
macro ' Rotate ';
procedure DoReslicing(horizontal:boolean);
macro '[H] Reslice Horizontally';
macro '[V] Reslice Vertically';
macro '[F12] Z Projection';
macro '[F14] Make Stereo Views';

LUT Manipulations
macro ' Invert LUT';
macro ' Log Tranform';
macro '[M] Gamma Tranform ';
macro ' Reset LUT';
macro ' Set Pixels Red ';
macro ' Nearly Gray LUT ';
macro '[U] Move Slice Up';
macro '[Y] Move Slice Down ';

Hot Keys

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6

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A

C

D

E

F

G

H

I

L

M

N -

O

P

Q

R

S

T

U

V

W

X

Y

F1

F2

F3

F4

F5

F6

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F8

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F11

F12

Macros