Setting up ‘Sensitized Emission’ based FRET experiments on Zeiss LSM780 confocal microscope

[Credits: Manoj V Mathew, Centre for Cellular and Molecular Platforms (CCAMP), Bangalore, India.
Experiments were conducted at the Central Imaging and Flow Cytometry Facility (CIFF), National Center for Biological Sciences (NCBS), Bangalore, India.]

Optical microscopes are resolution limited. However, there are ways to obtain sub-resolution information or measure sub-resolution distances in resolution limited images. One such technique is FRET.

FRET stands for Förster Resonance Energy Transfer. It involves two sets of molecules-one called 'Donor' and the other called 'Acceptor'. When a 'Donor-Acceptor' pair of molecules is in close proximity to each other the 'Donor' molecule can transfer its energy to the 'Acceptor' molecule. 

This happens non-radiatively via a dipole-dipole interaction mechanism. A 'Donor' molecule gets excited by excitation with its excitation wavelength. It transfers its energy to an 'Acceptor' molecule in its close proximity. The 'Donor' molecule then relaxes to the ground state without emitting a photon. The energy transfer results in the 'Acceptor'  molecule getting excited to its excited state and subsequently relaxing to its ground state radiatively (spontaneously emitting a photon).

These interactions are non-linearly (to the 6th power) dependent on the distance between the 'Donor' and the 'Acceptor'. They are significant only at very short distances between the 'Donor' and 'Acceptor', usually of the order of 10nm. By looking at FRET signals in diffraction-limited fluorescence images one can determine regions where the 'Donor-Acceptor' pairs are in very close proximity (<10nm) and measure the distance to an accuracy of the order of 1nm.

For FRET to occur there are some conditions the 'Donor' and 'Acceptor' molecules need to meet. For example, the 'Donor' emission spectrum should overlap with the absorption spectrum of the 'Acceptor'. One widely used FRET pair is ECFP and EYFP which satisfies this condition. Their spectra can be seen here.

There are many applications of FRET. One could study protein folding for example. If a FRET pair is tagged to the two ends of a protein, when it folds, the FRET pair gets closer to each other and exhibit stronger FRET signals. 

There are multiple ways to detect FRET. The following are some of them:

  1. Sensitized Emission: The 'Donor' is excited. The 'Donor' transfers energy to the 'Acceptor'. Acceptor fluorescence increases. 'Acceptor' fluorescence is measured.
  2. Acceptor Photobleaching: In a region of the sample, the Acceptor molecules are photobleached. This prevents the non-radiative transfer of energy from the Donor to the Acceptor. This results in an increase in 'Donor' fluorescence emission as more of the 'Donor' molecules in the region decays to the ground state radiatively. 'Donor' fluorescence before and after 'Acceptor' photobleaching is measured.
  3. Polarization Anisotropy: This technique is more useful for Homo-FRET ('Donor' and 'Acceptor' are the same species) measurements. Polarization anisotropy decreases with increasing FRET. Polarization anisotropy is measured.
  4. FRET by FLIM: This is probably the most sensitive FRET measurement technique. 'Donor' fluorescence lifetime decreases with increase in FRET. Donor fluorescence lifetime is measured most commonly using a Time-Correlated Single Photon Counting (TCSPC) based system.

For more details on the technique refer to the link

The idea of 'Sensitised Emission' would seem straightforward: Excite the 'Donor' and measure the fluorescence of the 'Acceptor'. However, there are few things we need to keep in mind. The very fact that we need the Donor's emission spectra to overlap with the absorption spectra of the Acceptor also means that there is inevitably some amount of:

  1. Cross Excitation: Some 'Acceptor' molecules will get excited directly by the 'Donor' excitation wavelength.
  2. Bleed Through: Some amount of 'Donor' emission will bleed into the 'Acceptor' channel.

This means that not all 'Acceptor' fluorescence is a result of FRET.  There are few other factors as well that can bias the results.

To have a more unbiased data from 'Sensitized Emission' based FRET experiments one needs to have a measure of these biasing factor. The following is the basic protocol:

a. Take three sets of images with the following sample sets:

  1. Sample with 'Donor' molecules only
  2. Sample with 'Acceptor' molecules only
  3. Sample with both 'Donor' and 'Acceptor' fluorophores

It is also possible to use only one sample if one can find regions within the sample with 1) only 'Donor',  2) only 'Acceptor' and 3) both 'Donor' and 'Acceptor'.

b. For each of these three sets take images using the following channels:

  1. Donor Channel: Excitation with 'Donor' excitation wavelength and collection in the 'Donor' emission spectrum.
  2. Acceptor Channel: Excitation with 'Acceptor' excitation wavelength and collection in the 'Acceptor' emission spectrum.
  3. FRET Channel: Excitation with 'Donor' excitation wavelength and collection in the 'Acceptor' emission spectrum.

In this write-up, I will describe how to use the Carl Zeiss LSM 780 confocal microscope to set up a 'Sensitized Emission' based FRET experiment using the 'ZEN' software.

Step 1: Setup the tracks and channels 

Setup three tracks with the following channels for sequential imaging with the following channels:

  1. Donor Channel
  2. Acceptor Channel
  3. FRET Channel

This is shown in Figure 1907/1.

Figure 1907/1: Setting up the three tracks 1) Donor Channel, 2) Acceptor Channel and 3) FRET channel

I used a fixed thin section of convallaria as the sample. While this is definitely not a good sample to demonstrate FRET, it is certainly enough for us to learn the software features. Convallaria is autofluorescent for excitations with most visible laser excitation sources. I used 488nm and 561nm excitation sources and spectral detector settings commonly used for Alexa 488 dye and Alexa 568 dye for the 'Donor' as well as 'Acceptor' and 'FRET' channels respectively. 

The channel configurations for the three tracks are shown in Figure 1907/2.

Figure 1907/2: Channel configurations for the three tracks- Left) Donor Channel, Middle) Acceptor Channel and Right) FRET channel

The following need to be kept in mind when configuring the channels:

  1. Use the same primary dichroic mirror for all channels
  2. Adjust the spectral detection bands to minimize bleed-through in the Donor and Acceptor channels.
  3. Use exactly the same spectral detection bands for both Acceptor and FRET channels

In addition, make sure that the Laser Power and PMT gain for each channel are set appropriately so as to avoid saturation of images. This is very important in any intensity quantification experiments. Saturation can be checked using the 'Range Indicator' feature in the software. It is also good to have the sequence mode as 'Change track every Frame' rather than 'Change track every Line'

Step 2: Acquire Images

Acquire three sets of images. Each set is imaged with one of the three samples using all the three tracks ( 1.only 'Donor',  2. only 'Acceptor' and 3. both 'Donor' and 'Acceptor'). This results in three image sets with 3 channels each (one each track).

In my case, I used a single sample and identified two regions 1) that had only 'Donor' and 2)only 'Acceptor' fluorescence. The whole field of view was used as the one with both 'Donor' and 'Acceptor'. For this, I first took an overview image with the track configuration described above. Figure 1907/3 shows the overview image.

Figure 1907/3: Overview image- Green: Donor Channel, Red: Acceptor Channel, and Blue: FRET channel

From the overview image, I identified regions with only green or red fluorescence. Using the 'Regions' feature the required region with only 'Donor' was marked for imaging. The 'Regions' setting is shown in Figure 1907/4.

Figure 1907/4: Regions setting

Make sure to uncheck bleach and check acquisition.

Check all the tracks and click 'Snap'. This will acquire an image of the ROI with only 'Donor' fluorophore in all the three channels. Save this image as 'Donor Only'

Similarly, use the 'Regions' feature to select a region with only 'Acceptor' fluorophores. Snap the image with all the three tracks and save the image as 'Acceptor Only'. 

The ROI images acquired using all the three tracks are shown in Figure 1907/5.

Figure 1907/5: ROI image with Donor Only (Left) and Acceptor Only (Right)

Now snap an image using all the three tracks of an ROI that has both 'Donor' and'Acceptor' fluorophores. I used the whole field of view. Call this image 'Donor and Acceptor'. In this case, the image will be same as the overview image I took. Now we have acquired three images each with three channels (one in each track). This is shown in Figure 1907/6.

Figure 1907/6: The three acquired images

Step 3: Set Donor and Acceptor parameters

Select the on the 'Donor Only' image and click on the 'FRET' tab on the side panel. This is shown in Figure 1907/7.

Figure 1907/7: FRET tab on the Visualization side panel

Note that the 'FRET' tab appears automatically under the following circumstances:

  1. Acquisition using 3 tracks (For 'Sensitized Emission')
  2. Acquisition using 2 tracks with one having a bleaching with time series (for 'Acceptor Photobleaching')

In the FRET GUI click on 'Parameters'. This is shown in Figure 1907/8.

Figure 1907/8: Setting Donor parameters

Do the following: 

  1. Select the Donor, Acceptor and FRET channels from the track assignments
  2. Click 'Donor' tab. This will calculate the Donor parameters Fd/Dd and Ad/Fd
  3. Select 'Donor' from the bottom drop down menu and click save. Save the parameters as 'Donor Parameters'

Similarly, select the 'Acceptor Only' image and click on the 'FRET' tab on the side panel. In the FRET GUI click on 'Parameters'. This is shown in Figure 1907/9.

Figure 1907/9: Setting Acceptor parameters

Do the following: 

  1. Select the Donor, Acceptor and FRET channels from the Track assignments
  2. Click 'Acceptor' tab. This will calculate the Acceptor parameters Fa/Aa and Da/Aa and Da/Fa.
  3. Select 'Acceptor' from the bottom drop down menu and click save. Save the parameters as 'Acceptor Parameters'

Kindly note that for the Convallaria sample I used has broad autofluorescence. The 'Donor' only and 'Acceptor' only regions may not exactly be that way. I selected them based on how they appeared in the overview image.

You can adjust the thresholds in the 'Thresholds' tab manually or select a background ROI in the image which will help the software calculate the thresholds. It would be advisable to define a background ROI (described below). The 'Thresholds' tab is shown in Figure 1907/10.

Figure 1907/10: 'Thresholds' Tab

For most applications set the 'Settings' tab to as shown in Figure 1907/11.

Figure 1907/11: 'Settings' Tab

Step 3: Analyze

Select the 'Donor and Acceptor' image and click on the 'Parameters' tab on the FRET GUI. Select 'Donor' from the drop-down menu. Load the 'Donor Parameters' file which was saved earlier. Then select 'Acceptor' from the drop-down menu and load 'Acceptor Parameters' file.

Subsequently, select the 'FRET' tab in the FRET GUI.  This is shown in Figure 1907/12. 

Figure 1907/12: 'FRET' tab

Select the Analysis method from the dropdown menu. There are three options available:

  1. Fc (Youvan)-"This method assumes that the signal recorded in the FRET channel is the sum of real FRET signal overlaid by donor crosstalk and acceptor signal induced by direct (donor) excitation. There is no correction for donor and acceptor concentration levels and as a result, the FRET values tend to be higher for areas with higher intensities". (Source: Zeiss LSM 880 ZEN 2 Manual)
  2. FRETN (Gordon) -"This method calculates a corrected FRET value and divides by concentration values for donor and acceptor. This method attempts to compensate for variances in fluorochrome concentrations but overdoes it. As a result cells with higher molecular concentrations report lower FRET values."(Source: Zeiss LSM 880 ZEN 2 Manual)
  3. N-FRET (Xia)-"This method is similar to the Gordon method with the difference that for concentration compensation the square root of donor and acceptor concentration is used. The resulting image is properly corrected for variances in the fluorochrome concentration." (Source: Zeiss LSM 880 ZEN 2 Manual)

Here I have used the N-FRET (Xia) method. 

Once a method is selected use the ROI tools in the FRET GUI to select a background ROI (a region with no signal). Check the 'Background' and 'Enabled' checkboxes for the background ROI. This helps the software to calculate the thresholds. 

Click the 'Analyze' button. This sted processes and analyzes the 'Donor and Acceptor' image to generate the FRET image. The FRET image will be displayed as shown in Figure 1907/13. 

Figure 1907/13: FRET image and FRET analysis results

Use the ROI tools in the FRET GUI to select an ROI on the image. The FRET analysis data for this ROI will be displayed below the image.

Kindly note again that I have used a highly autofluorescent sample which displays strong cross-excitation and bleedthrough. For the same reason, the calculated parameters might not be accurate enough as well. So please do not read too much into the FRET image and the results that are shown here. 

Performing FRAP experiments using Olympus FV3000 confocal microscope

[Credits:   Manoj V Mathew, Centre for Cellular and Molecular Platforms (CCAMP), Bangalore, India.
Experiments were conducted at the Central Imaging and Flow Cytometry Facility (CIFF), National Center for Biological Sciences (NCBS), Bangalore, India.]

FRAP stands for Fluorescence Recovery After Photobleaching. This is a very simple technique to study dynamics like diffusion rates in biological samples at the microscopic level. A better way to study dynamics would be at the single-molecule level. For such studies, FCS (Fluorescence Correlation Spectroscopy) might be a better tool. However, FCS is not easy, requires specialized hardware and some complex analysis modules. FRAP, on the other hand, can be easily performed on most confocal fluorescence microscopes. For a better understanding of the differences between FRAP and FCS kindly see (Link).

FRAP involves photobleaching a Region of Interest (ROI) in the biological sample and studying the recovery of fluorescence in that region over time. Since photobleached molecules cannot recover the ability to fluoresce, the only way recovery can happen is by the diffusion of non-bleached molecules from outside ROI. The average intensity of the ROI can be measured as a function of time and plotted as a graph. This can then be fitted onto standard diffusion models to compute parameters like diffusion rates. 

In this write-up, I will describe how to use the Olympus FV3000 confocal microscope for performing FRAP experiments.

The main interface of the FV3000 software is shown in Figure 1708/1.

Figure 1708/1: The main interface of the Olympus FV3000 confocal microscope software

Step 1: Image the sample

Use the software interface to generate an image of the sample. Adjust the laser power and PMT gain such that the image is not saturated but covers most of the dynamic range of the system. This can be ensured by using the HiLo mode in the image visualization window. I used a fixed thin section of convallaria as the sample. While this is definitely not a good sample to demonstrate FRAP, it is certainly enough for us to learn the software features. Convallaria is autofluorescent for excitations with most of the available visible laser excitation sources. I used 488nm excitation source and spectral detector settings commonly used for Alexa 488 dye for detection. 

In general, it is always recommended to use the 10% AOTF dampening (Laser ND filter set to 10%) to protect the detectors against excessive light exposure. This is especially important when using GaAsP detectors. However, when performing FRAP we need laser powers higher than 10%. Hence imaging should also be performed with the 10% AOTF dampening factor removed (Laser ND filter set to 'None'). In this case be extremely careful with the AOTF laser power setting and PMT gain setting to make sure you donot saturate the detectors.

Step 2: Enable the 'LSM Stimulation' and 'Synchronization' tabs

For performing FRAP experiments you need to enable the 'LSM Stimulation' and 'Synchronization' tabs in the software. Ths can be done by going to 'Tools Window' in the main menu of the software. This is shown in Figure 1708/2.

    Figure 1708/2: Enabling LSM Stimulation and Synchronization tabs from the Tools Window.

Once enabled they will appear within one of the tab containers in the interface as shown in Figure 1708/3.

Figure 1708/3: 'LSM Stimulation' and 'Synchronization' tabs added

Step 3: Set the 'LSM Stimulation' tab

Go to the 'LSM Stimulation' tab and set the photobleaching parameters. The 'LSM Stiumation' settings are shown in Figure 1708/4.

Figure 1708/4: 'LSM Stiumation' settings

The 'LSM Stimulation' tab provides the following:

  1. ROI tools: A number of tools to select one or more ROIs on the acquired image or the Live view
  2. Bleaching Dwell Time: The bleaching dwell time can be set different from the imaging dwell time. The image dwell time is entered in the 'LSM Imaging' tab. It might be useful to set a higher bleaching dwell time for effective bleaching.
  3. Bleaching Duration: You can set the bleaching duration to 'continuous'. In this case, the beaching continues till the user stops it. You can also enter a total bleach duration. For FRAP experiments it might be better not to use the continuous mode.
  4.  Bleaching Laser selection and powers: Bleaching is usually most effective with the same laser wavelength as the excitation wavelength of the fluorophore.  You would certainly need much more power for bleaching than what you use for imaging. But increasing the laser power much beyond the value where the fluorophore saturates is counterproductive. This is because at saturation, fluorophores achieve the maximum possible excitation-emission cycle rate. For photobleaching, most people simply blast the sample with maximum available laser power. This is not ideal. It would be great to perform a fluorophore saturation experiment first. Then set the bleaching laser power to about 10% more than what would saturate the fluorophore. After that play with the bleaching dwell time and total bleach duration for optimized bleaching. (Perform a few pilot bleach tests to optimize the parameters before commencing the actual experiment).

Step 4: Select one or more ROIs for bleaching and analysis

Once the "LSM Stimulation' parameters are set, select one or more ROIs on the image using the ROI tools in the 'LSM Stimulation' tab. Please note that selecting ROIs using the ROI tools on the visualization tab will not help.

I used the circle ROI tool to select a circular ROI. This is shown in Figure 1708/5.

Figure 1708/5: Selection of a single ROI on the image (labeled 1S)

Step 5: Set the Time Series

Set the Time Series from the 'Series' tab as shown in Figure 1708/6. 

Figure 1708/6: Setting the Time Series

The number of cycles and interval depend on how fast the fluorescence recovery is. If the recovery is fast, set the system to image as fast as possible ('Freerun'). It would also help to explore other ways of improving imaging speed like:

  1. Zooming into as small a regions as possible for imaging
  2. Decreasing imaging dwell time
  3. Decreasing the number of pixels (sometimes at the cost of resolution)

Please note that the total number of cycles set in the 'Time Series' also includes the pre-bleach cycles.

Step 6: Set the 'Synchronization' tab

Set the pre-bleach and post-bleach imaging cycles in the 'Synchronization' tab. This is shown in Figure 1708/7.

Figure 1708/7: Synchronization settings

Do the following:

  1. Base Method: Select 'LSM Imaging'
  2. Rest in stimulation: Select 'OFF'
  3. Stimulation Wait: Enter the number of required pre-bleach cycles. 


No: Post-Bleach cycles= Time Series Cycles - No: Pre-Bleach cycles

In my case: Time Series Cycles=30, Pre-Bleach Cycles=5, so Post-Bleach cycles=30-5=25.

Step 7: Run the photo-bleaching protocol

If only photo-bleaching is required and FRAP analysis is not required then go to the 'Normal' tab in the 'Aquire'' container and press 'Stimulation Start' as shown in Figure 1708/8.

Figure 1708/8: Photo-Bleaching only by pressing 'Stimulation Start' in 'Acquire' tab

Photobleaching will be performed on the selected ROIs as per the ROIs selected and settings in the 'LSM Stimulation' tab. A time series will not be run.

If FRAP analysis is required then, go to the 'Sync' tab in the 'Acquire' container and press 'Synchronization Start' as shown in Figure 1708/9.

Figure 1708/9: Sync Tab

The system will now run the time series: acquire the pre-bleach images, bleach the ROI/s and then acquire the post-bleach images as per the settings in the 'LSM Stimulation', 'Time Series' and 'Synchronization' tabs. The whole set will be saved as a time series. 

Note: Don't forget the mention the target folder for saving the 'Synchronization' files. This location is not mapped to be same as LSM acquisition ('Normal')  folder location.

The resultant image after bleaching and recovery is shown in Figure 1708/10.

Figure 1708/10: Resultant image post bleaching and recovery

Note: Since I used a fixed sample there is hardly any recovery (about 25 seconds post bleaching).

Step 8: Load the time lapse data into CellSens software for analysis

Open the acquired time series (with pre-beach and post-bleach) data in the CellSens software. This can either be done by opening the CellSens software and loading the file from the target folder or by right-clicking on the time lapse data on the acquisition software and clicking 'Export to CellSens'.

Once the time series data is opened in CellSens, we need to define two more ROIs in addition to the bleach ROI which is already defined and indicated in the time series data. The two additional ROIs are:

  1. Background: ROI in a region where there is no signal. This ROI would be used for background correction.
  2. Photobleaching Correction: ROI in a region which has signal but has not been bleached. This ROI will be used for correcting any bleaching that occurs during the image acquisition process after bleaching.

Note that the bleach ROI is already embedded in the time series data. But if you want to modify that or create a fresh one you can.

Make sure you select the appropriate ROI tools. This is shown in Figure 1708/11. 

Figure 1708/11: CellSens ROI selection tools for FRAP Analysis

Once the three ROIs (Bleach, Background, and Photo Bleaching Correction) are defined the time series data will look as shown in Figure 1708/12.

Figure 1708/12: Time series data with the 3 ROIs, Bleach Region (1S-Red), Background (ROI1-Blue) and Photobleaching Correction Region (ROI-2-Majenta) defined

Step 8: Plot the Intensity Profile

Click on the measurement tab on the top panel of CellSens. At the bottom of the tab, you will find tools for FRET and FRAP analysis. This is shown in Figure 1708/13.

Figure 1708/13: Tools for FRET and FRAP analysis

Click on the 'Intensity Profile' option. It presents a GUI shown in Figure 1708/14. Select the Time Series file location, Bleach ROI, and Background ROI and then press 'Execute'.

Figure 1708/14: Intensity profile GUI

This will display a background corrected intensity profile over time (both pre-bleach and post-bleach). This intensity profile as obatianed in my experiment shown in Figure 1708/15.

Figure 1708/15: Intensity profile over time

Note that you can choose to display the average, integral, maximum or minimum intensity on the intensity profile GUI. For the fixed sample I performed FRAP on, after bleaching while the fluorescence intensity drops very close to zero there is understandably no recovery of fluorescence over time.

Step 9: Perform FRAP analysis

You can now open the FRAP analysis GUI by clicking on the button on the top right corner of the intensity profile (Figure 1708/15) or use the option in the measurement tab (Figure 1708/13).

The FRAP analysis GUI is shown in Figure 1708/16. 

Figure 1708/15: FRAP Analysis GUI

As shown in Figure 1708/16  enter the following in the FRAP analysis GUI.

  1. Time Series file location
  2. Bleach ROI
  3. Single or double exponential fit. If the diffusion has only one component, use the 'Single Exponential Fit' and if the diffusion has two components select the 'DoubleExponential Fit'. For more complex diffusion scenario consider using some other fitting algorithm.
  4. Time series cycles over which the recovery data is fitted.
  5. Background ROI
  6. Photobleaching Correction ROI

The data fitting results are shown in the same GUI window. It also shows the raw and fitted data graphs. By pressing 'Execute' you can export the fitted data and results.


Hardware Configuration for Setting up Becker and Hickl Simple Tau 152 DX

[Credits:  Amit Cherian and Manoj V Mathew
Experiments were conducted at the Central Imaging and Flow Cytometry Facility (CIFF), National Center for Biological Sciences (NCBS), Bangalore, India.]

We have a Becker and Hickl (B&H) Simple Tau 152 DX TCSPC system. The system uses the two sets of detectors:      

  1. Two HPM-100-40 hybrid GaAsP detectors
  2. Two R3809U Multi Channel Plate (MCP) detectors. 

The detectors are attached to the Direct Coupling (DC) port of an LSM 780 Zeiss confocal microscope. The detectors are connected to the DC port using a A-Z-DC-710-PMZ beam splitter adapter from B&H. Having two detectors in each set is useful for techniques like Time Resolved Anisotropy (TRA), Fluorescence Lifetime Gated FCCS, or simply two color FLIM. A MaiTai Ti:Sa femtosecond laser at 80MHz is used for multiphoton excitation. 

The MCPs offer a shorter IRF but have a lower sensitivity and more after pulsing compared to the Hybrid GaAsP detectors. Depending on the application users prefer different sets of detectors. Hence we end-up swapping the detectors every once in a while. We thought it would be nice to describe the hardware configuration involved in connecting up the two sets of detectors, which is what we plan to do in this technical note.

Our B&H Simple Tau 152 DX TCSPC is a table top system that comes with a Laptop connected to a fast bus extension interface. The following cards are housed in the fast bus extension interface:  

  1. One DCC 100 detector control module 
  2. Two SPC 150 TCSPC modules (one for each detector)

Configuration 1:  Setup with R3809U Multi Channel Plate (MCP) detectors

Figure 1527/1 shows two MCP detectors connected to the DC port of the Zeiss LSM 780 confocal microscope using the B&H beam splitter. Note that mechanical shutter and the photodiode associated with each individual detector detectors were removed due to a small malfunction. The photodiode and shutter combination detect overload conditions and protect the MCP. If you are using the MCPs without this you need to be very careful about overload conditions. 

Figure 1527/1: Two R3809U MCP detectors connected to the DC port of Zeiss LSM 780 using the B&H A-Z-DC-710-PMZ beam splitter

The connection diagram for connecting two MCPs to the Sample Tau 152 system is described in the B&H TCSPC handbook and is shown in Figure 1527/2.

Figure 1527/2: Connection diagram for connecting two MCPs to the Sample Tau 152 system (From B&H TCSPC handbook)

The R3809U MCPs donot have internal high voltage generation circuits. Hence an external high voltage supply needs to be used. Our system uses a FUG HCN 14-3500 high voltage power supply as shown in Figure 1527/3. 

Figure 1527/3: FUG HCN 14-3500 high voltage power supply

The power supply system can supply upto 3.5kV. It has however only one output. Hence a voltage splitter is used to create two outputs to power the two MCP detectors. Figure 1527/4 shows the high voltage splitter. 

Figure 1527/4: High voltage splitter

This is a passive splitter. This is fine since high voltage supplies operate at very low currents. So splitting the voltage passively does to load the power supply.

The DCC 100 card has three connectors con1, con2 and con3. The con1 and con2 connectors are connected to the DCC1 and DCC2 connectors of the on the P Box. The P Box is powered by a 12 volts AC to DC wall mounted adapter. This is shown in Figure 1527/5. 

Figure 1527/5: P Box

The P Box has there outputs Shut1, Shut2 and Detect. 

Both Shut1 and Shut2 are connected to a connector with two output cables. These two cables are connected to the shutter and photodiode inputs respectively on the assembly of each of the MCP detectors. In our case since we had removed the shutter-photodiode assembly, we did not connect these. 

The Detect output of the P Box is connected to a connector with 3 cables. One cable is for the high voltage control of the power supply. This helps the power supply to be controlled and the voltage varied using the TCSPC software. In our case, we usually donot control the power supply using the software. We manually set the high voltage to a constant value of 3.4kV. The other 2 cables are for overload (OVLD) triggers and they are connected to the OVLD inputs of two widebad amplifiers.

The signal output of each MCP is connected to the IN pin of an individual wideband signal amplifier (B&H HFAC-26dB) as shown in Figure 1527/6. 

Figure 1527/6: B&H Wideband signal amplifiers

The amplifiers are powered by a 12V AC to DC wall mounted adapter. The signal output of each of the two wideband amplifiers is connected to the Constant Fraction Discriminator (CFD) input on the two individual SPC-150 TCSPC boards housed in the fast bus extension interface.

A breakout box BOB-4 is used to split the scan-clock from the LSM-780 confocal microscopes onto the two SPc-15 cards. Figure 1527/7 shows the breakout box.

Figure 1527/7: B&H Breakout box

The scan input of the breakout box is connected to a connector with 2 cables. The two cables are connected to the scan clock outputs at the back of the Zeiss LSM 780 real time controller. This is shown in Figure 1527/8. If the breakout box is used only for splitting the scan clocks, it need not be externally powered. 

Figure 1527/8: Scan Clock connections on LSM 780.

The SPC-1 and SPC-2 outputs are connected to the scan clock inputs of each of the SPC-150 cards.

The Sync inputs for each of the two SPC-150 cards are obtained by bleeding a small amount of the femotsecond laser light at 80MHz using a thin glass slide onto a B&H PHD-400 high-speed photodetector. This is shown in Figure 1527/9.

Figure 1527/9: Generating Sync input using PHD-400 high-speed photodetector

The output of the photodetector is split using an SMA Y splitter as shown in Figure 1527/10 and fed to the SYNC inputs of the two SPC-150 cards.

Figure 1527/10: SMA Splitter

It should be made sure that length of the two cables and effective delay in them of the two coaxial cables connecting the SYNC inputs of the two SPC-150 cards as similar as possible. Also, the length should be adjusted to get the optimal timing. Else additional patch cords should be added.

Configuration 2:  Setup with HPM-100-40 hybrid GaAsP detectors

Figure 1527/11 shows the connection diagram for connecting two HPMs to the Sample Tau 152 system as described in the B&H TCSPC Handbook.

Figure 1527/11: The connection diagram for connecting two HPMs to the Sample Tau 152 system (From the B&H TCSPC Handbook).

The setup is very similar to the one described above for MCP detectors. The main differences are:

  1. HPMs have internal high voltage generators. Hence an external high voltage source is not required.
  2. The Con1 and Con3  connectors on the DCC card are used.
  3. P Box is not used
  4. Con1 and Con3 connectors on the DCC are directly connected to the power supply and control inputs on HPM1 and HPM2 respectively.
  5. The HPMs have internal broadband amplifiers hence the HPM signal outputs can be directly connected to the CFD inputs on the respective SPC-150 cards.