Zeiss LSM 410 confocal manual


The Zeiss 410 is the Comprehensive Cancer Center's shared use confocal, which is maintained and supported by the Light Microscopy Core Facility. The instrument is a robust imaging system based on the Zeiss Axiovert 100 inverted microscope. UV, green, red and far-red laser lines are available and high-quality images of up to three fluorophores can be produced simultaneously.

LMCF staff are here to help you - we will train you to use the instrument and will provide technical support and imaging assistance as needed. If you have any queries or difficulties, please contact us at LMCF@duke.edu or 613-8168.

Conventions used in this manual

  • File menu options and subheadings are listed in sequence in boldface type: File→ Store Image
  • Field names & slider scales are written in bold typeface.
  • Italics are used to note common problems and helpful tips.
  • Window titles & manual section titles are written in quotations: "Create New Database" 

Words of precaution

Using oil immersion objectives
  • Use the MINIMUM AMOUNT of immersion oil.
  • If you are looking at a number of slides under oil immersion, you may find it necessary to wipe oil off the objective (with lens paper or cotton swabs) every couple of slides or so.
  • At the end of your session, wipe off oil from the immersion objectives with lens paper or cotton swabs.
  • Do not try to use the 10x or 20x objectives after using oil on a slide. The optics will be terrible and you will almost certainly get oil on the non-immersion objective. It is OK to use the 5x objective. If you get oil on a dry objective, please let Heather (613-8168) know immediately and she will clean the objective.

If you fail to follow these instructions, there is significant risk of oil seeping down into the objectives and causing expensive damage to the optics of the microscope!!

Risk of damage when focusing on specimen

If you ever feel ANY resistance when turning the focus knob (whether going up or down) STOP TURNING THE FOCUS IMMEDIATELY! Because of the mechanical advantage in the coarse adjust, it takes very little force to damage the objectives and/or the focus control gearing.

The top of 40x or 63x objectives should be even with the top of the stage plate when those objectives are in focus. Set this first by eye before you put your slide on the stage, then change to the low power objective that you wish to use, place your slide on the stage, and adjust focus using ONLY the fine focus control

If you follow this simple procedure, you won't damage to the microscope, the objectives, or your specimen. More complicated preparations (like multiwell plates) require more complicated procedures and greater care on the part of the user. PLEASE see Heather or Sam if you plan on using an unusual setup.

System Start Up
  1. Sign into the logbook, listing the date, your name, your PI's name, and the start time of your imaging session.
  2. Close the fluorescence shutter and put the reflector slider in the "confocal" position by pulling both all the way to the right. 
  3. If the system was left running by the last user, skip to step 7 or 8 below.
  4. If desired, turn on the mercury lamp for conventional fluorescence (non-confocal) microscopy. The power supply is located on top of the air table, to the left and just behind the microscope stand.
  5. Turn on the red/green laser power supply (the riser between the two monitors) by turning the key switch to start and releasing it (just like starting a car). You should hear the fan turn on automatically. After ~30 sec, there should be a subtle click when the laser actually ignites. If clicking continues, turn laser off and report problem.
  6. Turn the computer/microscope/far-red laser using the key switch on the cabinet to your right just beneath the air table (also starts like a car ignition switch).
  7. If using the UV laser, turn it on now according to "Using the UV laser" near the end of this user manual.
  8. Once the operating system loads, you will be asked for a user name and password to log on to the ASADMIN network domain. In the log-on window, make sure the domain field says ASADMIN and not confocal or some other domain name. Once this information is entered, click on the LSM icon on the desktop to open the software. 

Conventional Microscopy

Put slide on stage oriented perpendicular to front edge of table with its coverslip down as in Figure 1. The coverslip should fit entirely within the hole in the center of the stage (use a 22x30 mm or smaller coverslip). You may add a small piece of tape at one end of the slide to secure it, but this usually is not necessary.

Most objectives are designed to use #1.5 coverslips. Using the wrong one may have serious implications on image quality. Please refer to the link for "coverslip thickness" which can be found on the LMCF website.

Fig 1) Proper Orientation Of Slide On Stage

  1. Put the reflector slider into position 2 by moving it one click-stop to left. This is the UV/transmitted light filter for viewing your specimen with transmitted light. Refer to Figure 2.
  2. Fig 2) Diagram of Confocal Mirror and Fluorescence Filters In Reflector Slider

  3. In the Control Panel, click the toggle button <> between Conv and LSM modes once.
  4. Select the T radio button for transmitted light under Conv. Turn on the halogen lamp using the rocker button on the power supply box located to the left of the microscope stand. Adjust brightness as needed by using the dial on the power supply.
  5. Start with a low power objective - either 5x or 10x. The condenser (above the stage) should be in the "DIC .5-1.4" position. Focus the specimen and center it in the field of view using counter pressure on the 4 knobs that project up from the floating stage. Refer to Figure 3.
  6. If you have trouble finding your specimen, close the condenser diaphragm all the way (small lever on front of condenser all the way to the right). This will improve contrast and make your specimen easier to find.

    Fig 3) Condenser and Floating Stage

  7. To switch to an oil immersion objective (40x, 63x), rotate the objective nosepiece (by gripping the knurled edge of the nosepiece, not the objective) until the desired objective is just to one side of the slide (see figure 4). Place a very small drop of oil onto the front lens of the objective.
  8. Avoid actually touching the objective with the applicator. Avoid getting bubbles in the oil by letting the oil flow into the nozzle by gravity for a moment before squeezing the bottle very gently.

    Continue rotating the nosepiece until the objective clicks into position beneath the slide.

    Fig 4) Applying Oil To the Lens

      Reminder: once you have used an oil immersion objective on a slide you cannot use the 10 or 20X dry objectives with that slide. In fact, you must use caution when changing objectives to avoid contaminating the dry objectives.

  9. Carefully refocus the specimen (fine focus only for high mag objectives!!) and re-center if necessary. If you wish, you may change its orientation in the field using the circular stage. This stage is just below the floating stage and can be rotated by its knurled edge. Refer to Figure 3. Assuming proper stage centration, your specimen should stay in view as you rotate the stage. Do not use the knobs that project upward from the floating stage for rotating the specimen.
  10. To observe fluorescence, click R or FL under Conv in the Control Panel. You may wish to turn off the transmitted light power supply at this time if it is no longer needed. Move the reflector slider to a position appropriate for the fluorochrome used (see Fig 1).
  11. Open the fluorescence shutter (push left one position). You should now see fluorescence through the eyepieces. To avoid unnecessary photobleaching, it is good practice to close the shutter (see Fig 5) whenever you're not looking at your specimen in the eyepieces.

Fig 5) Fluorescence Shutter Closed & Open

Single channel confocal image acquisition

1. Close the fluorescence shutter (slide fully to the right). Move the reflector slider all the way to the right to the "confocal mirror" position (you may hear a very short beep).

2. Click the toggle button <> between Conv and LSM in the "Control Panel".

3. Open the fluorescence shutter (slide over one position to the left).

4. Check the red-green laser power output by clicking POWER above the "Control Panel" near top of screen. The power level should be set to 30 for most applications. If using UV laser, set its power level according to the UV instructions near the end of this manual.

5.Under the Lens= menu at the top of the "Control Panel", select the objective you are using.

The Lens setting is very important! Since there is no electronic communication between the computer and the nosepiece, all calculations made by the software (scale bars, z projections, etc) will be derived from this information you provide.

6. Under the At= (attenuation) menu, you will find 3 columns of numbers (1, 10, 30, etc). This menu selects neutral density filters for the laser light entering the microscope. The left column controls the far-red laser (for Cy5, Alexa633), the middle column controls attenuation of the UV laser, and the right column controls the red/green laser. Start with the far-red and red/green lasers at 10 (=1/10 laser light) and the UV laser at 3. If you find you need more excitation power, then set the attenuation to a lower number.

7. Select the proper dichroic mirror by putting the lever on the front panel of the scanning unit marked DBS2 in the "Free" position. This is as per Table 1 (the "Cheat Sheet" describing the dichroic positions for single, double, and triple channel imaging combinations) which is included in this manual and posted in the imaging suite.

Table 1) Dichroic Mirrors and Emission Filters Cheat Sheet

8. For green-labeled specimens, click radio button 1 in the Setup box on left side of the "Control Panel". For red imaging, click 2. For far-red, click 3.

9. In the Scan box, click Start. You should see a flickering light at the tip of the objective. If not, check the position of the fluorescence shutter (step 3).

10. Adjust focus with the fine focus knob until you see the specimen on the display monitor. You may see red & blue pixels in the image - this is the range indicator color table (Image→ Color Tables→ Range Indicator if it appears you are not in this mode). Range indicator is used in setting brightness & contrast of the image. Adjust focus until you see the brightest image, which will probably have many red pixels in the areas where signal is detected.

11. Press F9 on the keyboard to obtain initial brightness (offset) and contrast (gain) settings. Optimize these settings by adjusting the Contrast scroll bar (click and drag slider with the right mouse button for fine adjust) so that about 10% of the pixels are red (these will be the brightest ones in the final image). Adjust the Brightness scroll bar so that the background is no more than 50% speckled with blue pixels where you do not expect signal (these will be the darkest pixels in your final image).

12. Recheck focus. If you find that the image becomes much brighter, then readjust contrast and brightness as before according to this brighter focal plane. Click Stop when not actively adjusting the image to avoid bleaching your specimen.

13. To improve image quality, select "Averag" in the Extras box of the "Control Panel". Select line averaging and the number of iterations you would like the laser to scan each line. Click Done in the Averaging window.

The amount of averaging necessary to obtain a good image depends on the signal to noise ratio of your sample image. Low contrast images will require more averaging but be aware that this could cause photobleaching. You can determine the amount of averaging necessary to yield the best image of your sample empirically, but most samples will require averaging of 4 or 8.

14. To collect the averaged image, click Single in the Scan box.

15. For a gray scale display, click Color in the "Display Control" window 3 times or select Image→ Color Tables→ gray scale from the main menu. For a colorized display, click the RGB radio button in the Image box of the "Display Control".

16. To save the image, choose File→ Store Image. You should see the contents of the Bio-lemming\Confocal server, which has been mapped to the H: drive. (The FROG confocal server is mapped to the F: drive.) Scroll down to your lab's folder, then double-click to open. Repeat as necessary to enter your personal user folder. Clear the contents of the File box and type a name (up to 8 characters) using lower case letters and/or numbers only. Click Store to save image. Files are saved as a Tiff, which can be read on either PC or Macintosh computers.

You can use Windows to make new folders with longer names describing your experiment - ask us if you do not know how to do this.

17. To increase magnification of a scan:

    a. Click the small unlabeled button to the right of the Zoom scroll bar at the bottom of the Control Panel. The Zoom and Offset window will appear.
    b. To set zoom, left-click on Set below the red box. A green outline will appear on the display monitor.
    c. Move the position of this defined region by left-clicking the inside of the region and dragging. Change the size of the region (smaller box=higher magnification) by left-clicking and dragging a corner of the box region.
    d. To exit the display monitor, click the right mouse button once.
    e. Left-click Done in the Zoom &Offset window. Note that the minimum zoom (Zoom min button) equals 0.8.

18. To switch back to conventional microscopy:

    a. Close fluorescence shutter.
    b. Slide the reflector slider to the left(over two notches for the green filter).
    c. Click the toggle button <>between Conv and LSM.
    d. Reopen the fluorescence shutter to observe through the eyepieces.

Dual or triple channel confocal image acquisition

  1. Select Param→ Load User Parameters from the main menu. Double-click the parameter that matches the combination of fluorochromes you are using.
  2. Because of the 8 character limit to the length of file names, the pre-set parameters for general use are named after the cyanin fluorochromes with UV fluors referred to as "dapi" or "blu". Therefore, it is important to know that Cy2 is a green fluor, Cy3 is a red fluor, Cy5 is a far-red fluor. Considering this naming convention, "-cy2cy3" is the setting for double-channel red/green imaging and "-c2c3blu" is the triple-channel setting for imaging the green, red, and UV fluor combination. These are all explained in the cheat sheet.

  3. Upon loading a user parameter, the computer will set everything on the microscope except the DBS2 and DBS3 mirrors, which are manual. Consult the chart just below the monitors (hanging from the shelf) to find the correct positions of these mirrors for the combination of fluorochromes you are using.
  4. Contrast (gain) and brightness (offset) are set separately for each channel. To select a channel, click the appropriate color under Image on the Display Control window. For example, for Cy2/Cy3 double labeling, click the R radio button to adjust the red channel. Now adjust contrast and brightness for the red channel as before (step 11 above). To adjust the green channel, click the G radio button, and then adjust contrast and brightness again. Far-red signal is displayed in the blue channel when triple channel imaging.
  5. To display multiple channels simultaneously, click RGB in the Image box of the "Display Control".
  6. To save the images, use the same procedure as in step 16 of "Single channel confocal image acquisition". The system saves the image AS IT IS DISPLAYED at the time you click Store. To save all channels, make sure that RGB is clicked. To save just one channel in gray scale, select the appropriate channel (R, G, or B) and then save file.

Double- or triple-channel imaging using the SEQUEN function

In some cases (e.g. severe bleedthrough conditions) it is better to sequentially acquire various color channels of data from a sample. This can be done manually, by exciting each fluorochrome individually and saving the images to different color channels of the video monitor. The various color channels are overlaid by clicking RGB in the "Display Control", and the composite image can be saved in the usual manner (File→ Store Image).

The SEQUEN and SEQUEN3 functions automate the sequential excitation, acquisition, and display overlay of this process. Their use is described here.

  1. The "Control Panel" must be set-up for single-channel excitation as it is at system start-up. In this set-up, a user can sequentially excite green, red, and if desired far-red, in exactly that order. If you need to excite some other color combination besides green-red or green-red-far red, let Heather know and she can write a custom user parameter for your imaging scenario.
  2. Set your brightness and contrast for each color channel in your sample. Make sure to change the display monitor's color table from range indicator (red & blue pixels) to greyscale after doing so.
  3. Set your Averaging as desired.
  4. If you're imaging just two color channels, press SEQUEN. If you want to image three channels of data, press SEQUEN3.
  5. SEQUEN will acquire every channel and will load the overlaid images in the display monitor. Store this final image to your folder on the server, if desired.

Fig 6) Location of SEQUEN Buttons In Software


1. After setting the brightness and contrast for every channel in your specimen, determine the start and end points of the Z scan. Z sectioning is always done moving into the specimen, away from the coverslip. To find the starting point for the Z series, Start scanning (with the averaging off) and focus toward the coverslip (rotate fine focus knob toward you) until you reach the point at which you'd like to begin the Z series. Stop scanning.

2. Select Z→ Initialize motor from the main menu. The Z value in the "Parameters" window should now read 0.0.

3. Start scanning. Turn fine focus knob away from you (counterclockwise) focusing down through your specimen until you reach the point at which you'd like to end the Z series. Stop scanning and note the Z value at this point in the specimen.

4. Manually focus back to the Z=0.0 position. Set averaging to your desired value.

5. Select Z→ Z Sectioning under the main menu. Set your desired Z interval (optimal settings depend on the pinhole size, the objective and the wavelength of the light but as a starting point try 1 µm for 40x objective, 0.5 µm for 63x objective). Set the number of sections to cover the interval between the start and finish points of the series:

    number of sections needed = (thickness of sample / z interval) + 1

Set current section position to 1.

6. Specify where the Z series files will be stored. Select File in the Destination box. Click Dir that appears to the right to select the folder to which the series will be saved. Select your folder on the server (H: drive). Type in an alphanumeric file name up to 6 characters long (do not end with a number). Click Select. The name you provided should now appear in the box just to the left of Dir.

7. Click OK to begin Z sectioning. Each image of the series will be stored as an individual tiff file with numbers (starting with 00) appended to the name you specified.

Z-Sectioning using the DUALSCAN macro

The DUALSCAN macro sequentially excites & acquires multiple fluorochromes in a single specimen at fixed intervals over a specified distance. Currently, this macro is set up to collect only two color channels of information from a sample.

DUALSCAN works by exciting the first fluorochrome, collecting its emission signal, and writing that data to a designated color channel in video memory. The macro then excites the second fluorochrome, collects its emission, and writes it to a contrasting color channel of video memory. Finally, it displays the overlay of the two collected images before moving on to the next user-defined z-interval. The user can specify to save the monochrome image series of each color channel and/or that of the RGB overlaid images. Sequentially excited images are saved to the directory as they are acquired.

  1. The "Control Panel" must be set-up for single-channelexcitation as it is at system start-up. In this set-up, a user can sequentially excite green and then red, in exactly that order. If you need to excite some other color combination besides green-red, let Heather know and she can write a custom user parameter for your imaging scenario.
  2. Set your brightness and contrast for each color channel. Make sure to change the display monitor's color table from range indicator (red & blue pixels) to greyscale after doing so.
  3. Determine the depth of interest to be z-scanned by following Steps 1-3 in "Z-sectioning".
  4. Manually focus back to the Z=0.0 position. This is critical!! The DUALSCAN macro is practically incapable of focusing to the top of a z-stack before starting the image series acquisition.
  5. Set Averaging to your desired value.
  6. Select Macros→ DUALSCAN from the main menu.
  7. Once in "Dualscan", press Get LSM Set-up. The macro records the acquisition settings for Set-ups 1 and 2 in the "Control Panel".
  8. Fig 7) Dualscan Window

  9. Press Edit Z-scan Parameters. In the "Z Sectioning" window that opens, designate your desired z interval. In the Number of Sections field, set the number of sections necessary to cover the interval between the start and finish points of the z-stack:
  10. number of sections needed = (thickness of sample / z interval) + 1

    Fig 8) Z Sectioning Window

  11. Set Current Section Pos. to 1.
  12. Select the Save checkbox beside 1st Series Files if you want to save the monochrome image series collected in Set-up 1.
  13. Click Dir_1 to the right of the checkbox. In the window that appears, designate the directory path to which the series of images will be saved (ie your folder on the server). Highlight everything except for the ".tif" in the field at the top and give the z series a root name of no more than 5 alphanumeric characters long. Press Select in the bottom left corner of the window. Press Return to return to the "Z-Sectioning" main window.
  14. Fig 9) Directory Path Selection Window

  15. Repeat the process of directory path selection and file naming for the 2nd Series Files if you wish to save the monochrome image series collected in Set-up 2. The Save checkbox must be selected in order for this file series to be saved.
  16. Repeat the process from Step 10 in order to save the RGB Series of composite (overlaid) images.
  17. Press Done to return to the DUALSCAN macro's main window.
  18. Press Z-scan. In the window that opens, you should see details of all of the z-sectioning information programmed while in "Z-Sectioning". Press Ok to start the z-series acquisition.
  19. Pressing Break will stop the dualscan image acquisition, and the focus drive (z motor) returns to the top of the z Images taken to that point in the acquisition will have been saved on the specified directory path.

  20. When the scan series is complete, click Exit. DUALSCAN will close and a window will appear that says, "Data: DSC are modified! Save changes?" Select No.

Ending your session

1. Sign out on the logbook by listing end time. Indicate whether or not the system was left on for the next user. Note any problems in the Comments column of the logbook.

2. Select File→ Quit from the main menu or close the program using the X in the upper right hand corner of the main window.

3. 3. Check the on-line reservation schedule. (Use the Internet Explorer icon on the 410’s desktop.) If someone else is scheduled to use the microscope within 2 hours, then select the "Log Off" command from the Start menu of the desktop and leave everything running. The log in window should appear. However, if it is near the end of the day or on the weekend, please contact the next person on the schedule to ensure that they are actually planning to use the microscope. If you can't contact them, then shut off the system.

4. If you have been using the UV laser and no one else is signed up to use the UV in the next 2 hours, turn it off now. Leave the chiller running to cool the UV laser down for the remainder of the shut down process (at least 3 minutes). Refer to the "Using the UV laser" for full details of UV laser power down.

To turn off the system, select "Shut down computer" command in the Start menu of the desktop. Wait for a message stating "It is now safe to turn off the computer" to appear. Turn off the computer by turning the key on the cabinet below the air table to the off position.

6. Turn the key switch on the KrAr laser (on shelf above the microscope) to the off position. The fan will continue to run for several minutes then will shut off automatically.

7. Turn off the mercury lamp (on shelf above the microscope).

8. To make cleaning the microscope stage and objectives easier, gently tilt the upright condenser stand back out of the way.

9. Clean oil immersion objectives by gently removing the excess oil from the objective surface with lens paper and/or a cotton-tipped swab. Do not use any solvent! The non-immersion objectives (5x, 10x, 20x) should not be cleaned by users of the facility. If you note oil on a non-immersion objective then notify LMCF at lmcf@duke.edu.

10. Rotate the objective holder by its knurled edge (not by objective) to put the 5x or 10x objective in place.

11. If the stage and/or benchtop surfaces have oil on them, wipe them clean using a kimwipe dampened with alcohol.

12. Return the condenser stand to vertical position.

13. Cover the microscope with the blue cover. If the mercury lamphouse is still hot, cover everything except the lamphouse.

14. Turn off the UV chiller switch if necessary.

15. Turn off the room lights and close the door behind you (unlocked). Make sure the main door to the Imaging Suite (4226) is locked behind you when you leave. This door should always be kept locked when nobody is there.

16. When you return to your lab, download your files onto some other storage media (e.g. your department's fileserver, memory stick, CD/DVD) right away. If you do not know how to do this, then please seek help from your department’s computer support personnel. Files will remain on the server for 1 week, but after that time are subject to deletion without notice.

Using the UV laser

General guidelines for UV usage

  • If you intend to use the UV laser, indicate this in the Comments field when booking your reservation on-line so other UV users will be aware that you are using the laser that day
  • Turning the UV laser on & off throughout the course of the day should be kept to a minimum. For this reason, there is a "2-hour Rule" for the UV laser:
    If someone is signed up to use the UV within 2 hours after your UV session, leave the UV laser on for that user. If the next UV user is scheduled to start more than 2 hours after your UV session, turn the UV laser off according to the directions below.
  • The computer and other lasers do not have to be off or shut down before turning on the UV laser.

Turning the UV laser ON/OFF

  1. The water-driven chilling unit provides cooling water for the laser and is located in the equipment closet, room 4229. Turn the chiller on by using its main power switch located on the unit. This white rocker switch requires a firm compression into the “ON” position in order for it to actually stay on.
  2. Turn the UV laser on via the UV power supply, the black box that sits on the floor to the right of the isolation table. Flip the main power breaker switch located above the key switch to the "ON" position. Now turn the key switch to the "ON" position to start the laser.
  3. In ~20 seconds the UV laser will ignite. There will be a short "scream" at this time - this is normal. The CRT monitor display may look scrambled. Regardless of how it looks to you, it should be degaussed by pressing the button labeled "Degauss" located on the face of the monitor below the screen.

Preparing to image with the UV laser

  1. You can now log into the Zeiss software, if you haven't already. If you want to image UV staining alone, call up the single channel detection preset for UV via selecting Parameters→ Load User Parameters from the main menu and double-clicking the "-dapi" setting. Put the DBS2 mirror goes in the "Open" position as it would with any other type of single channel detection.

    There are preset imaging combinations containing UV (double and triple channel) available for use, as well. As usual, reference the chart hanging from the shelf below the monitors to find the correct positions of the D2 & D3 mirrors for the combination of fluorochromes you want to image.
  2. Before imaging your sample, you will need to set the laser power outputs using POWER2 located outside of the "Control Panel" window, several buttons to the right of the POWER button that you're used to using. Once in the POWER2 window, "Power 1" controls the red/green laser and should be set to 30 as usual. "Power 2" controls the UV laser with 50 being a reasonable starting value. You may have to adjust this power level as your sample dictates.
  3. The UV laser requires a 20-30 minute warm-up period for optimal excitation of your specimen.

  4. When finished, consult the on-line reservation schedule to determine if you need to turn off the UV laser according to the "2 hour Rule" as stated in the "General guidelines for UV usage". If you need to turn off the UV laser, turn the key switch on the UV power supply to the "OFF" position, then flip the main power breaker "OFF". After having completely turned off the power supply, wait 3 MINUTES before turning the chiller off to allow proper cool down of the UV laser.
  5. Finally, indicate in the UV ON/OFF column of the log book whether or not you turned the UV laser off. Don't forget to reference the ON/OFF status of the rest of the system.

Hints for better UV imaging

  • The 40x is the best objective to use when imaging UV signal. Being a Plan NeoFluar objective (which is not quite as highly-corrected as the 63x), the 40x is optimized for high light transmission. The 63x objective is a Plan Apochromat objective, which has lens elements that don't transmit UV as well. You will notice significantly decreased image quality (e.g. low UV signal, noise, accentuated “hot spots”) if you image UV with the 63x.
  • The following have been saved to all of the UV imaging configurations but should be kept in mind in case you write your own UV configuration or if you make adjustments to the pre-set UV imaging parameters:
    - When imaging any UV fluorophore, scan with both UV laser lines (351nm & 364nm) simultaneously. This improves the signal from the specimen.
    - Don’t use any Attenuation on the UV laser. Check that the middle column of the At drop-down menu at the top of the "Control Panel" is set to “1”.

Image sequence applications

All of these features require that a sequence of files be preloaded into the host memory (or RAM) of the computer.

  1. Select File→ Load Sequence from the main menu.
  2. Direct the files you're loading to the computer's RAM by clicking on Host Memory box located underneath the Format box on the right side of this window.
  3. Select the sequence you wish to load from the list of files appearing on the left. Highlight on any one image file in the series and either double-click or press Load to load the entire sequence of files. It may take a moment for the entire series of files to load.


  1. To view the series, select Z→ Animate from the main menu. The box next to "From host" should be selected. Also select Reverse.
  2. Click Start to begin paging through the series. If you wish to view just one channel, click the appropriate color at the bottom right of window.
  3. Clicking Stop will stop the animation. Click Cancel to exit.


  1. Select Z→ Gallery from the main menu.
  2. In the Number of Sections field, enter any number > actual number of files in the loaded series. In the Source box, activate the Memory radio button. Select the Draw Frame box, if desired. Select either the Display Data or the Display File Name boxes, if desired.
  3. Click Ok. The series of files is displayed on the image monitor and can be saved as a tiff image via File→ Store Image.
3-D maximal projection

The projection software creates 3-dimensional views using alpha blending as a model for reconstruction. It is easiest to think of alpha blending as illuminating the specimen from different angles or viewpoints.

A maximal projection is one which takes the intensity values from all sections and collapses them into one single illuminated image. "Maximum" overlay is selected when the maximum pixel intensity is desired to be represented in the projection. "Transparency" overlay mode decreases the brightness of each file in the projected stack to reveal structure in subsequent layers that would otherwise be hidden. Creating a maximal projection is described here.

  1. Select 3D→ Projection from the main menu.
  2. In the Mode box activate the y Axis radio button. In the Overlay box activate the Maximum radio button. Leave the Angle field at 0 and the Center field at 50 if an en face (straight-on) projection is desired.
  3. Click Calculate. An image with virtually infinite depth of focus is calculated and displayed on the image monitor.
Producing scale bars on saved images

The Zeiss software can provide scale bars for any confocal image. For these to be accurate however, you must have selected the correct objective under the lens menu (see step 5 under the "Single channel confocal image acquisition") before you collected and saved the image. Normally the scale bar is written into a channel called the overlay channel. Although it appears on the screen, it is not written into a standard part of a tiff file and therefore does not appear on the screen when a file is exported into another program such as Photoshop.

There are two methods for transferring the bar from the overlay channel into one of the regular channels so that it can be exported. Either will work on the confocal microscope computer itself, while only the latter will work on the workstation computer.

Method I (Confocal microscope computer only)

This method uses a macro called OVLTORGB (overlay to RGB), which is located near the top of the Zeiss confocal software menu. This macro converts the normal scale bar (held in a part of memory called the overlay channel) into a line that is written onto the image. To produce a scale bar:

  1. Select Function→ Measure, and click the Scale Bar checkbox. Clicking the Set Bar button allows you to position the scale bar (position by moving mouse). Exit this mode by clicking the left mouse button.
  2. Close this window - the scale bar should remain on the image.
  3. Click OVLTORGB to write the scale bar into one channel of the image.

Because the scale bar has jagged edges, you will probably want to save this modified file under a different name - once the OVLTORGB command is performed you cannot remove the scale bar from the image. You can then use this file as a guide for making a new neater looking scale bar on the original image using Photoshop, Powerpoint or another program.

Method II (Confocal microscope or workstation computer)

Method II uses the measure function to provide crosshairs on the image at a defined distance that can then be used to produce a scale bar. This method will work on either computer.

  1. Select Function→ Measure from the main menu, and click Posit in the measure window.
  2. Position the two crosshairs anywhere on the screen (click and drag each with left mouse button). The number display shows the distance in microns between the two crosshairs.
  3. Position the crosshairs at a convenient place and distance for a scale bar (eg. just below the distance display).
  4. Click on the measure window to exit positioning mode.
  5. To write the crosshairs and distance display into the red channel of the image, click on Draw Graphics and Draw Display.
  6. Use File→ Store image to save a copy of the file under a different file name. When you open this image in another program, you should now find the crosshairs and distance reading. Use this to create a scale bar for the original image as described in Method I.