Frequently Asked Questions (FAQ)

For the ICTM method: none! However, this is only true when the a-priory knowledge that the object is non-negative is absolutely true, as is the case for fluorescence emission.

Yes. Huygens Essential treats the image as the only known plane of a 3D stack and proceeds as usual. Set the z-sampling distance to the Nyquist rate as explained in 'sampling densities' (Huygens User Guide).

The deconvolution process enhances the fine structure of the cells, but unfortunately if the image contains artefacts (because of unstable laser power etc.) they will also become more apparent in the deconvolved image. Scanner instability is mainly a not-reproducible phenomenon, thus deconvolution is not the solution. In some cases the instability IS reproducible like slow thermal drift. If this instability is also present in the PSF measurement the deconvolution can reconstruct the image.

The absolute value of the final Quality factor much depends on the data, the microscope type, and the background. It is a global value computed over the entire image, so the contribution of a local resolution increase can be small.

For example, suppose you have a large featureless image with one tiny object. While the tiny object may be restored very well, the change in the featureless part is negligible. The quality factor will therefore hardly change, though the restoration is successful.

Usually, widefield images show a much higher quality increase than for example confocal images.

TIRF (Total Internal Reflection Fluorescence) deals mostly with 2D images. Huygens can handle 2D images by internally treating them as part of a 3D stack from which most planes happen to be missing.

Strictly speaking Huygens does not generate TIRF Theoretical PSFs, but customers report good results with high NA confocal PSFs. Proceed by setting the Microscope type of your image to confocal and NA ~ 1.4.

An experimental PSF, if existing, can naturally be used. See also the wiki article Total Internal Reflection.

Yes, Huygens will certainly improve the image.

The first test should be done using a Wide Field Microscope Theoretical Psf with the actual parameters used for the acquisition, maybe with a lower Numerical Aperture to enlarge the PSF a little bit.

Still, remember that deconvolution assumes that the Image Formation is linear, and transmission is not, due to possible interference effects. These effects are lesser for thicker samples, but they can create restoration artifacts if they are noticeable. These can be balanced by properly tuning the Signal To Noise Ratio and the Max Num Of Iterations.

The ideal particle to distill an Experimental Psf with the Psf Distiller is probably a sub-resolution gold particle. Note that all the images should be inverted (negative) in Huygens before distilling a PSF and deconvolution: high intensities (gray values closer to white) should describe high object density, but raw transmission images provide the opposite. In Huygens Essential, this can be done with Tools > Invert image.

The Ideal Sampling constraints are about half as stringent as for the confocal imaging mode. For an objective with high N.A. (1.4) the voxel sizes should be in the range of 100-150 nm laterally and 350-500 nm axially.

This strongly depends on the refractive indexes (lens immersion medium and specimen embedding medium). If there is no refractive index mismatch then restoration up to 200nm can be expected.

The restoration improves the resolution in all directions, but more so in the z-direction. In typical confocal images it is easy to increase the z-resolution by a factor 2. With a measured PSF a factor 4 is attainable.

However, the z-resolution in the measured image is often 4x worse than the lateral resolution. So at best you can compensate for that, but due to the lateral resolution gain the result will still be non-spherical. The gain in lateral resolution can be spoiled by applying a Gaussian filter (Operation window -> Restoration -> Gaussian filter/Quick Gaussian filter) to decrease lateral resolution again in a controlled manner. Because, in our experience, practically no one wishes to reduce resolution the restoration tools do not do that automatically.

Still, if the sphere is physically small enough (of the order of a voxel) the restoration can reduce it to close to a single voxel. Provided that the z-sampling is small enough. This situation occurs easily in the deconvolution of widefield images with 100nm x 100nm x 100nm voxel size.

For fluorescence and from a theoretical viewpoint MLE restoration is the only proper choice. However, for high SNR's as in widefield images this becomes rather academic. Since the ICTM method in Huygens Pro is computationally more efficient than the MLE implementation we suggest using ICTM for WF images. As to 'quality', an examination based on subjective criteria such as visual inspection might lead to ICTM winning in low noise cases; an examination based on mathematically sound criteria has shown MLE to be the best choice. For noisy images (most confocal ones) the MLE algorithm is not only scientifically more correct, it also produces visually more pleasing images: far less background noise artifacts than ICTM.

The computation time depends on a large number of factors:

• Microscope type: WF microscopes require more iterations than confocal or 2-photon microscopes.
• Object type: sparse objects can be restored more effectively than dense objects. The more gain is possible, the more iterations are needed, even if the iterations themselves become also more effective ('bigger steps').
• Noise: low noise makes a large gain possible: more iterations are needed.
• Algorithm: Our ICTM (Iterative Constrained Tikhonov-Miller) iterations take per iteration less time than our MLE (Max Likelihood Estimation) algorithm.
• Hardware: faster and more processors speed up things, insufficient memory is problematic: processing speed then depends on disk I/O performance.
  • Example: Restoring a moderately noisy confocal 256x256x64 image, including starting the software, loading the image, generation of a PSF and a MLE run take altogether 4:23 minutes on an SGI Octane with 2xR10000@225MHz

The deconvolution itself is done frame by frame, but the pre and post processing need the whole time sequence. The preprocessing consists of bleaching correction and background estimation. In Huygens Pro there is also the possibility to apply a time or full 4D prefilter. In the postprocessing the Z-drift is corrected over time (see Zdrift Correction). Most of these operations work on multiple adjacent frames, so working on a file by files bases would limit the current and future processing possibilities.

This does not imply that the 4D stack image needs to be loaded in RAM memory. Provided the swap space is large enough most of it can be swapped out. In fact, paging out means that the processed data is written in raw form back to disk. This will slow down the preprocessing operations, but the deconvolution speed would hardly be impeded, paging being only necessary between frames.

A second possibility is to use a script to process the frames individually. Because the CMLE and QMLE deconvolution methods allow you to specify a background level as a percentage of the estimated background that would handle varying backgrounds also nicely.

There can be two major reasons for this:

• There is a problem with the measured PSF:
  • bead images were saturated or undersampled.
  • beads have formed aggregates.
  • beads were moving while being imaged.
  • insufficient signal from beads causing inaccuracies in the averaging procedure.
  • strongly varying background.
• The conditions under which the PSF was measured do not match the imaging conditions.
  • The most important parameter is the medium refractive index. To exclude magnification calibration problems, best record the bead images at the same sampling density as the specimen. To check whether there is a matching problem between the measured PSF and the specimen data, deconvolve the specimen with a theoretical PSF. If the result is better then there can be a matching problem.

As a preliminary check on the PSF quality, proceed to deconvolve one or more of the bead images with it. This should result in a strong gain in resolution.

When you deconvolve a 2D-time image as a 3D image, you force the software to assume that the relationships between the 2D slices, if any, are due to the axial imaging properties of the microscope, but this is obviously not the case! The results will be therefore incorrect.

Instead, when you correctly process the 2D-Time Series as such with the Time option the software will, after correcting for bleaching and variable background, properly deconvolve each 2D image as a 3D data set with one plane recorded and the rest missing, as explained here Is deconvolution on 2D or 2D-time images possible?.

Some File Formats having indexes in the file names are interpreted as 3D stacks by default: you may need to convert the dataset from XYZ to XYT once opened.

As an alternative you could write a Tcl script for Huygens Scripting or Huygens Professional to deconvolve the 2D-time series frame by frame. Still, your images would not be corrected for bleaching and varying background, a task automatically performed if you have the Time option.

In case of a 3D-time series, with the Time option your data will be also corrected for axial drift.

Yes. The results are especially remarkable for Widefield Microscopes. A 2D image recorded with a microscope can be considered as a slice from a 3D image. In this case the Huygens Software treats the data as a (severely) truncated 3D image, but 3D nonetheless. When deconvolving it Huygens attempts to reconstruct blur sources outside the slice and remove blur from it, regardless of the time frames.

Proper parameters

When doing 2D deconvolution of widefield images, set the Z Sample Size to the ideal Nyquist value. You can calculate this by using the Nyquist Calculator.

If you have a 2D time series, make sure Huygens interprets the series as such. Some File Formats having indexes in the file name are interpreted as 3D stacks by default: you may need to convert the dataset once opened from XYZ to XYT. See Convert The Data Set. The restoration might not be optimal otherwise, as explained in Can a 2D-time series be deconvolved as 3D stack?.

Yes, this is certainly possible in Huygens.

This can be done in Huygens Essential by choosing the option 'invert image' from the 'Tools' menu before starting the deconvolution. In Huygens Professional select the image and go to 'Deconvolution', 'Operations window', 'Arithmetic', 'One image', and then 'Invert'.

Then proceed to deconvolve the image. For brightfield images we strongly advice to use the linear Tikhonov Miller algorithm (available in Huygens Professional) as this algorithm does not amplify the background noise.

Brightfield imaging is not a 'linear imaging' process. In a linear imaging process the image formation can be described as the linear convolution of the object distribution and the point spread function (PSF), hence the name deconvolution for the reverse process. So in principle one cannot apply deconvolution based on linear imaging to non linear imaging modes like brightfield and reflection. Fortunately, in the brightfield case the detected light is to a significant degree incoherent. Because in that case there are few phase relations the image formation process is largely governed by the addition of intensities, especially if one is dealing with a high contrast image, 'linearizing' the problem. In short, a Bright Field Microscope is not exactly a linear imaging device, but can be made to behave almost like one.

Dr. Marcel Oberlaender et al. from the MPI of Neurobiology in Martinsried proved the validity of the Huygens deconvolution for brightfield data with the linear Tikhonov Miller algorithm.

In practice one goes about deconvolving brightfield images by inverting them and processing them further as incoherent fluorescence widefield images. Still, one should watch out for interference patterns (periodic rings and fringes around objects) in the measured image. These could become pronounced in low contrast images.

Using the Operations Window in Huygens Professional all channels are processed in one run. To work with only one channel proceed to split the multichannel image (Select Edit->Split or ALT-S) into single channel images. The single channel images can be deconvolved individually with specific parameter values. For example, the SNR can be edited in the "Signal/Noise per channel" input line of the Operations Window. The Join operation can be used after the deconvolution of single channel images to combine them into a multichannel image.


If using the Devonvolution Wizard of Huygens Professional or Essential, you can choose which channel you want to process and skip those you are not interested in.

Besides the Huygens deconvolution FAQ there is plenty of information on image acquisition and restoration, deconvolution, and the way the Huygens Software works available at the SVI site. Below, a number of links to help you get started:

• Deconvolution
• Image formation
• Restoration parameters
• Deconvolution algorithms
• Huygens manuals

It can be rigorously proven that when dealing with non-negative objects (fluorescing objects) the I-divergence criterion (MLE) is the only consistent choice, whereas for objects which can be both positive and negative (e.g. sound) least squares (ICTM) is the best choice. Another viewpoint is that I-divergence incorporates the Poisson nature of the emitted fluorescence light whereas least squares incorporates Gaussian noise. With this in mind one could expect that for low noise levels where the differences in the Poisson and Gaussian distributions are small there is no preference between the methods other than their computational efficiency. One reason why we still have a preference for the MLE algorithm is that it handles noisy backgrounds much better than the ICTM. A disadvantage of MLE is that it easily overemphasizes small structures, but we constrain this.
See also: MLE versus ICTM - Will one method be faster or more rigorous or give a better quality result?.

As a rule of thumb one gets easily 2x in Z and a bit less in XY. This applies to even noisy images. To gain more resolution a measured PSF is necessary. Some published figures (Bioimaging 4 1996 pp. 187-197): HIW (nm) z x y

• raw bead image 790 270 265
• restored 221 116 93
• bead object function 83 83 83 (i.e. the `true' bead)
• difference 138 33 10

Conclusion: nearly 4x in Z, more than 2x in XY.

The quality factor (QF) can only be used in a relative way, to compare between iterations, and is used for the stop criterion. The comparison of these values makes no sense if applied to deconvolutions of different images. The Quality Factor of the MLE algorithm is directly derived from the I-divergence; in the ICTM algorithm it is derived from the Tikhonov-Miller functional as described in the literature.

Sometimes one channel of the dataset contains Reflected light signal, i.e., the light which passes straight through various beam splitters and on into the Reflection Detector.

1. Is there any validity in deconvolving this Reflected light component?
2. If so what emission / excitation wavelength's would be applicable?

This is very tricky since reflected light is coherent and full of interference effects. On top of that quite some microscopes have serious trouble with interference between the signal and stray light. In short, deconvolving reflection images is not a good idea. If you still want to proceed, set the excitation and emission values to the wavelength of the reflected light.

Yes. Single plane widefield deconvolution works because the data is extrapolated into a region above and below the plane spanning typically between 10-20 planes of 100-300 nm in Z. The software generates an appropriate PSF.

The QMLE iterations are approximately 5 times more efficient than the CMLE iterations, whereas they also take slightly less time per iteration. So 10 QMLE iterations are equivalent to 50 iterations in CMLE. CMLE is superior in handling low Signal to Noise (SNR) data, like low light level confocal images. In principle the CMLE algorithm with an SNR setting > 60 converges to the same result as the QMLE algorithm, but after many more iterations. Also, for good quality widefield images QMLE is the best choice.

See Restoration Methods.

The deconvolution process increases the Dynamic Range of the dataset, i.e, the intensity range increases. A computer screen cannot display high intensities arbitrarily, there is a maximum for the brightness of a pixel in the screen. Therefore the maximum intensity in an image is normally mapped to the maximum intensity a screen can display, and all the other pixels in the image are scaled accordingly. Read more here.

People have done simulations with synthetic objects:

• van der Voort, H.T.M and K.C. Strasters, "Restoration of confocal images for quantitative image analysis" JoMi 178, 1995, pp 165-181.
• van Kempen, G.M.P. et al, "Comparing Maximum Likelihood Estimation and Constrained Tikhonov-Miller Restoration". IEEE Eng. in Med. And Biology 15 No 1. pp 76-83, 1996.

The voxel sizes are not changed by the restoration! The slicer takes the exact aspect ratio into account. No rounding off to integers.

Usually, when this happens you have run out of swap space. If you work with large images it is better to switch off the undo system. As a rule of thumb, bear in mind that 3 or 4 times the size of the float image can be reserved by Huygens out of the available RAM memory. The rest can be swapped out. When working with large images in Huygens then the disk I/O speed is an additional bottleneck. As a result, the time needed to swap pages in and out during an iteration determines computing speed.

Consider a 300MB dataset (~25Mvoxel image): the system has to write AND read *at least* 300MB per iteration. To speed up swapping it is advantageous to count on fast disks. Additionally, the current ICTM algorithm cannot process large datasets brick by brick. The CMLE and QMLE methods both can process datasets brick by brick. We recommend to use CMLE in general and QMLE in particular for low noise widefield data.

We have a set of batch scripts available on our webserver: See Batch Script. Notes:

1. The script is able to launch multiple parallel jobs to make full usage of a multiple cpu system.
2. These scripts use Huygens Scripting (license needed). Please ask for a 30 day license (info@svi.nl) if you'd like to test this. You can add the license to your system using the Help > License > Add License tool.

See also Batch Processor.

The multiphoton feature can be set up in the 'Excitation photon count' field in the Microscopic Parameter Editor. For a 2-photon system set this field to '2'. See Multi Photon Microscope.

These steps will help to lower the amount of memory needed for deconvolution:

1. Switch off the undo system (Options menu).
2. Reduce the size of the image (Crop tool). In particular, widefield images often contain many slices with blur.
3. A theoretical PSF will be generated on-the-fly by the MLE and QMLE restoration methods. There's no need to create a theoretical PSF by hand.

When you convolve two images the resulting intensity is spread over the corners of the image. This phenomenon is caused by the following two properties of the FFT:

• Images are interpreted as periodic, i.e. the image has infinite size but repeats itself in each dimension with periods that are equal to the size of the original image in the corresponding dimension.
• The frequency origin (frequency space) is located at the zero voxel (bottom-left-plane) of the 3D image. The positive frequencies are located in the first octant of the image. Depending on the type of transform you selected, `complex' or `real', the negative frequencies are present in the other octants.

This means that you will find spatial frequency 0,0,0 at voxel (0,0,0) of the transform, and not at the center. For visualization purposes it is often desirable to have the zero frequency centered. Use To/from optic rep. from the Restoration menu in the Operations Window to move the zero frequency to the center.

`Real' Fourier transforms contain only the positive frequencies in the u-direction (with u, v, w , spatial frequencies corresponding with the x, y, z axis). However, they do contain negative frequencies in the other dimensions. For visualization purposes real-FFTs are therefore less suited than complex-FFTs.

As a consequence of defining the frequency origin (frequency space) at the (0,0,0) voxel, convolutions with funtions which are not centered around (0,0,0) will cause a shift in the resulting image. For example, when you convolve an image with a sphere located at the center ( xc , yc , zc ) of the second image, the result will be shifted over a vector ( xc , yc , zc ). Because images are interpreted as periodic all octants in the resulting image will appear `swapped'. You can prevent this from happening by using the following methods:

• Center the image with which you are going to convolve around (0,0,0). When this image, for instance a sphere generated with Generate sphere , is centered use To/from optic rep. to shift it to the origin. In other cases use the following method:
• (Alternative centering method) Determine the Center of Mass (CM) of the second image with Image statistics. Then use Shift image to move the CM to the origin. Since Shift image also interprets the image as periodic, this will produce the desired result. An advantage of this method is that you can shift the CM over a non-integer distance.
• When you have convolved with a function located at the center of the image, you can undo the shifting effect by applying To/from optic rep. to the convolution result.

Yes, key publications are:

Csiszar, I., 1991; Why least squares and maximum entropy? An axiomatic approach to interference for linear inverse problems. Ann. Stat., 19, No. 4, pp. 2033-2066 (PDF 8 MB).

Kempen, G.M.P., van der Voort, H.T.M., 1006; Comparing Maximum Likelihood Estimation and Contrained Tikhonov-Miller Restoration. IEEE Engineering in Medicine and Biology vol 15, No 1, pp. 76-83

Manufacturers supply the average size along with a standard deviation. Should we look for very tightly specified beads?

The spheres we developed and tested this technique with, had a coefficient of variation of 2% at 230nm diameter. This variation was smaller that the precision at which we could locate the center of the beads, at best 1/4 pixel. So if the c.v. is a good deal better than say 1/4 of a pixel it is good enough.

Around 30 optical units — an optical unit is (for NA=1.3 and lambda = 500nm) 61nm — so about 2 micron. Because the software doesn't know whether there is a bead just over the edge of the image, the bead-edge rejection distance is the same.

If the 175 nm beads yield enough signal best use them.

During the "Reconstruct psf from bead image" phase in Huygens Professional, or during the PSF distillation in the PSF Distiller, the question "center of mass of the image containing the averaged bead is far (0.482 microns) from the center of the image - proceed anyway" popped up.

This is a check in the Reconstruct PSF tool to prevent the cones (fans) of WF PSFs to be cut off too asymmetrically. Also, this effect can be caused by large spherical aberration making the PSF (and therefore the bead images) highly asymmetrical. This check was built in the software to make people aware of sub-optimal properties of their data which should be corrected during acquisition.

The averaging procedure can increase the signal of the accumulated bead as much as needed. BUT it makes an alignment error which is worse when signal is poor. The alignment error effectively increases the bead size, but not in a way which can be predicted easily. As soon as the bead signal is sufficient for alignment the SNR can be increased by averaging, but if it is insufficient you're sunk. We've never seen enough signal from 50nm beads. On *very* sensitive microscopes 110nm beads are good enough, but we doubt whether they are safe to use on regular commercial microscopes.

For more information you can have a look at DeconvolvingBeads

Several restoration operations can optionally center the channels of the PSF. The option "Center PSF" aligns the center of mass of the channels of the PSF. A non-centered PSF will cause a shift of the restoration result. This can be advantageous to correct the image for chromatic shifts, but we advice to use the Chromatic Shift Corrector for that after deconvolution.

See also Color Shift.

Each image has a microscope type associated to the microscope (sensor) with which it was taken. Currently confocal, spinning disc, widefield, multiple photon, 4 PI and STED images are supported by Huygens. By default Huygens does not know the sensor type and assigns "Generic". However, the method for generating a PSF is different for each of the microscope types. There is no particular method to generate PSFs for generic (unknown) microscope types.

To assign a microscope type to the image:

• select the image
• choose "Edit Parameters" or press Alt-P
• change the field "Microscope Type" from "Generic" to the correct microscope type
• save the changes.

A PSF can now be generated based on the image.

The message about the axial criterium may mean that the beads are too close to the top and bottom edges. In that case a bead below or above might disrupt the PSF. Since usually the beads are adherent to the glass there is little danger of this so you can ignore it.

But it may also mean that the image is indeed having not enough information about the PSF along the optical axis Z. The recorded volume around beads, specially in widefield microscopes, must be large enough to containg information about the light cone. If this is the case, better image the beads again, maybe changing the sampling density and recording more planes. Read more at Recording Beads.

No. The reason here is that the shape of the expected (theoretical) PSF is forced only lightly on the measured PSF. This is done in the form of a bandlimit constraint, and the pinhole size has not much influence on this. If we would impose the theoretical shape on the measured data more strongly it would, for instance, force a non-existent axial symmetry on the measured PSF, exactly what we don't want. Still, from version 2.07 of the Huygens onward the theoretical shape is imposed on the data slightly more strongly.

Yes. Since the wavelength parameter has an effect on the PSF you need to generate or measure a PSF for each different channel. The deconvolution wizard handles multi-channel images for you and the PSF distiller also detect multi-channel beadimages to generate a multi-channel measured PSF. In scripting you process a multi-channel image by first splitting it into a series of one-channel images. Then you deconvolve these images using the measured or generated PSF's for each channel and merge the results.

To obtain a measured PSF the Huygens Software contains a set of tools with which you can derive a PSF from images of fluorescent beads. In principle one would need a very small, point-like bead to measure a 'point' spread function. Since sufficiently small beads are also very dim this is impractical. Huygens solves this by being able to extract PSFs from larger, much brighter beads. A typical size is 200nm radius. To improve accuracy it is also capable of averaging over multiple beads, from single or multiple images.

See also Psf Distiller.

It all greatly depends on the Numerical Aperture (NA), to a lesser degree on WaveLengths. For a 'typical' Confocal Microscope (NA = 1.3) 5 micron is fine, 3 will do. For a Wide Field Microscope the situation is different because the PSF does not stop in Z. For restoration the software needs a PSF which is a bit larger in Z than the image to be restored. Now the problem is that this needed extent can't be known to the software during the measurement, i.e. bead averaging/cleaning + PSF reconstruction.

It turns out that in practice recorded PSFs are often too shallow, so we have fitted out the reconstruct tool with a powerful extrapolator and a manual Z-size setting. Provided the extrapolator has a big enough foothold it can extrapolate to whatever size you need. Rule of thumb for the foothold is 5 micron. Ideally the extrapolator is called as part of the preprocessing phase of the restoration tools, but because it is rather lengthy we put it in the reconstructor. We did put a light-weight extrapolator in the restoration tools to be able to extend a PSFs a little bit.

See also Recording Beads.

Here are some interesting links:

• Summary of Molecular Probes' fluorescent beads can be found through this link (microspheres)
• List of Polysciences' fluorescent beads can be found through this link
• List of Thermo Scientific particles

Beads with a size close to the width of the PSF half intensity width (HIW, the width of the PSF at 50% of the peak intensity) show up in the image as spots which are slightly wider than the PSF. To compare a bead image to a theoretical PSF proceed as follows:

• Open an operation window on the bead or averaged bead image
• Choose a free destination image
• Click the PSF button and set Dimensions to Parent
• Click Run to generate a theoretical PSF of which the size matches the size of the bead image

You can now compare the two images by opening a Slicer window on the PSF destination image. If you find that the beads are much larger than the PSF you might be looking at bead clusters. In that case you might find weak single bead images in the background. Because the clusters are of unknown size you can't use them for PSF measurement. Alternatively there might be large beads in the sample. If these have an unknown size, or if their size is larger than the HIW, best not use them for PSF measurement.

Using the Nyquist calculation I calculate that the minimum step in Z should be 170nm (x40water, NA 1.15, ex 364nm). I see from the example in the manual that you sampled a 230nm bead at 25nm steps.

If you like you can oversample the beads at say 2x the Nyquist rate in the lateral direction. In general it is best to really match the Nyquist criterion (or better) in z since highest resolution gain is in Z. It also depends on the capabilities of the z-stepper. If you use finer 170nm beads instead of 230nm beads (and get sufficient signal), so much the better. It has no impact on the sampling density. Of course there is a relation between the Nyquist rate and the bead size, but it is a weak one. Literature: van der Voort HTM and Strasters KC (1995) Restoration of confocal images for quantitative image analysis. J.Micr. Vol 178, pp 43-54.

The deconvolution tools are capable of adapting a PSF derived from such bead images to the sampling density of the specimen. Still, it is best to sample at the same density as at which you are going to collect the biological images later, or with densities which differ by a factor of 2 or 3.

In principle, it is sufficient to record bead images for a PSF calculation at the system Nyquist rate, say 50 nm × 50 nm × 150 nm for a confocal setup (or doubled figures for widefield). If your sample of interest has been imaged at a different sampling rate, then the PSF can be adjusted by bandlimited interpolation. This is automatically done by the Huygens CMLE and QMLE deconvolution methods. You can use our Nyquist Calculator for determining the ideal sampling values for your experimental conditions.

There are two problems, though:

• In some microscopes the magnification at high zoom is unreliable, with errors up to 30%.
• In case of widefield data, the Nyquist sampled PSF might be too small (in terms of microns) to be used in deconvolving physically large data. Make sure you specify, during the PSF distillation, a large enough required size for the final PSF. However, in extreme cases memory limits might get in the way.

If you want to obtain confocal-like 3D images from regular epifluorescence microscopy, i.e. widefield microscopy, then you need indeed a series of images recorded at different depth: a so-called stack. To arrive at a result that is comparable to a well-sampled confocal image a typical Z step size (Z sampling distance) is 200nm.
If you'd like to obtain confocal-like single slice images the best procedure is to acquire a short stack of 10-20 slices around the plane of interest and deconvolve that. However, if you lack a z-drive or the time to acquire the stack Huygens-Pro and Huygens Essential also allow you to deconvolve a single 2D widefield image.

Basically it is very simple: beads were brought on a coverslip and left to dry. Then for water type medium simply distilled water was added. For high refractive index Aquatex (Merck) was added, covered with a second coverslip and left to dry for 3-4 weeks. For this last sample 115 nm beads from Molecular probes were used with very good results. For a recipe using glycerol see section "Measuring a PSF" in the recipe booklet that you received in the case you have purchased the software.

A 1micron bead is too large for a PSF measurement because it lacks sufficient high spatial frequency components. In other words: a large bead has a lower surface/volume ratio than a small bead. Therefore it has reduced 'edge energy' per unit of total signal strength. In addition, beads as large as 1 mu are often not stained homogeneously and are likely to distort the PSF due to their high refractive index. All these factors render 1 micron beads unsuitable for PSF measurement.

One could argue that very small beads(<25nm) have ideal spectral content, but up to now such small objects lack signal strength. Averaging small beads doesn't work either since the limited signal strength limits the precision of alignment procedure. This situation might change when quantum dots become available for PSF measurement.

The alignment procedure in the Huygens System is based on determination of the Center of Mass (CM) of the beads. This allows bead alignment with sub-pixel accuracy. We found that with 150-230nm diameter beads the right balance is struck between spectral content and alignment accuracy. However, when the fluorescence yield of beads can be increased by better dyes or better anti bleaching agents the optimal size will be reduced. Good results have also been obtained with 110nm beads.

Measuring both the PSF and the sample with the Ideal Sampling is always nice, but not strictly necessary. If required, the deconvolution functions in the Huygens Software will automatically scale the measured PSF to adapt it to the image sampling. However, the PSF should always be ideally sampled or better. In order to do deconvolution, a slight undersampling can occur in the sample image but only to a certain extent.

Why is the Nyquist Rate sampling so relevant for deconvolution? The (degrading) imaging process acts at the scale of the PSF, and therefore this must be precisely acquired in order to restore the image properly. See Ideal Sampling for more details. Therefore, in any case, the beads for PSF acquisition should be imaged with a Sampling Density at least according to the Nyquist Rate, or even better. Like that the PSF would contain all the information about the imaging properties of the microscope, and can be adapted to other imaging conditions that are slightly undersampled. See also the FAQ What is the maximal voxel size at which Huygens can still do a good job?.

PSF measurement

The Huygens Software will reject beads that are severely undersampled if you try to distill a PSF from them, because in that case they do not contain the necessary information to do it! The Nyquist Rate is the minimum sampling required for a proper PSF measurement. Oversampling the bead image can be a good idea (it increases the Signal To Noise Ratio of this fundamental image), but in practice this is not possible in the widefield case, because the image would be too large. Because other microscope's PSF are smaller, you can afford some oversampling there. If possible limit the differences in sampling density to factors 2 or 3, thus making the later scaling of the PSF easier and more precise.

In practice (and with good signal) it is not necessary to sample finer than 25 nm lateral and 100 nm axial for confocal systems or 50 nm lateral and 100 nm axial for widefield systems. Fair numbers in a typical confocal case are 50 nm lateral and 150 nm axial.

Caveat: at high zoom factors the magnification as reported by the microscope is not always reliable.

In the widefield case best record with no Pixel Binning. This usually results in a lateral sampling density in the 67-100 nm range. Axial sampling should match the sampling of the specimen if it is below 250 nm.

For more information see Recording Beads and Parameter Variation.

Indeed, the better your PSF, the better the resolution to which you can deconvole the object and the fewer artifacts. But as to the resolution, there is a limit. Signal To Noise Ratio (SNR) plays a crucial role there.

The Nyquist Rate (similar to the Shannon theorem) says that IF a signal is bandlimited (see our FAQ What's a bandlimited system?), it is sufficient to sample it at twice the highest frequency. Then, it is possible to reconstruct the signal at ALL locations, perfectly. So in principle it is sufficient to sample at the Nyquist rate. Taking more samples does not get you more information about the object. In short, the ideal sampling rate is not infinite. Still, taking more samples with the same number of photons per pixel will improve the quality of the deconvolution result. Vice versa, taking more samples allows you to achieve the same quality in the deconvolution result at lower photon counts per pixel. BTW: If you sample below the Nyquist rate you get Aliasing Artifacts (moire patterns, straircasing).

One more reason to oversample is that with sparse objects and good SNR it is often possible to achieve a Half Intensity Width resolution on the objects corresponding with a Band Width in excess of the microscope's bandwidth. The objects are then said to be super resolved. The Shannon theorem says it doesn't matter whether you get the supersampled image during sampling or afterwards by interpolation, but it is more practical to get it during sampling, if only to improve the SNR situation.

A different matter is two-point Spatial Resolution: separating two objects. It is very hard to separate two objects reliably at distances smaller than the Nyquist distance.

To see what is the ideal sampling for your setup see Nyquist Calculator.

No, not necessary!

If the PSF is not centered deconvolution will also shift the image. Usually this is unwanted, so the PSF generator and current PSF measurement tools always center the PSF. In previous versions of Huygens this was not always the case, so for example the ICTM has an option to center the PSF.

In multi-channel images the different channels are often shifted with respect to each other. By NOT centering the PSF this shift can be automatically undone by the deconvoluttion, and with sub-pixel accuracy. On the other hand, the shift can be just as well done manually with by the shift tool, also with sub-pixel accuracy. This is perhaps the most practical method.

Since Huygens 3.7 there is a specific option available that measures and corrects for shifts between channels. See Chromatic Shift Corrector.

With large beads the software has a much harder job extracting a PSF than from images of small beads. However, small beads are noisier and will need some averaging to reach a sufficient Signal to Noise Ratio (SNR). Very small low signal beads can't be averaged because their position cannot be measured with sufficient accuracy. As a rule 175 nm diameter beads yield sufficient signal while being small enough. We have seen good results with 110nm beads, but we do not think these can be imaged with sufficient SNR by all microscopes. For high NA microscopes (NA > 1.2) we do not recommend beads as large as 260nm.

When generating a PSF some choice must be made about the size. In most of the cases a border is added to avoid edge artifacts (padding). Depending on the purpose a choice can be made.

1. Ideal: The size (in microns) of the generated PSF is determined from the physical size. Since the fringes of a PSF go on forever it is in theory not spatially limited, but in practice a volume can be chosen beyond which there is a negligible amount of energy from the PSF. The exception is the widefield PSF: there the amount of energy outside a volume around the focus is always infinite... As the intensity goes down locally it is possible to find a point at which the intensity of the PSF is well below the accuracy of any camera. Still, ideal WF PSFs are much larger than confocal or 2-photon ones.
2. Parent/Padded Parent/Full Padded Parent: The size is derived from the Parent image: either exactly the same as the parent in fact no padding is made here, or as large as if the parent was 'padded'. The extra volume computed by the software is a trade-off between FFT (Fast Fourier Transform) compute efficiency and the size of the original image. For example If you have 31 layers in your image, adding one layer would optimize the Fourier Transform process. But adding one layer is not enough to prevent wrap around effects. The software will find out how many layers extra is a good compromise. The Fully padded parent mode is relevant for widefield images, for other microscope types this is equivalent to Padded Parent. If PSFs are to be compared it is best to use 'Parent' because that will fix the size.
3. Automatic: A tradeoff is made between the physical size of the PSF and the memory requirement. In practice confocal or 2-photon ones are ideally sized; WF PSFs are smaller than ideal but at least as large as the padded parent.
4. Manual: You can manual set the number of Z-slices in this mode using the input field "Min XY-slices (Manual)". Widefield images should never be padded manually.

Full question: We tried to reconstruct PSFs from beads taken in wide-field mode. The problem is that Huygens excludes all the beads from the calculation because of different reasons: too close to an edge or too close to each other. Even taking 100 slices did not improve the situation. We know that the image of the bead in widefield mode can be extremely large even if the cut-off is high.

Yes, this is the problem. Even worse: the thicker you make the stack the wider they become so the tops of the cones will tangle. That will really mess up the PSF. Best way to go is to set 'reduce PSF size', for instance to 2 (=high), and reduce the number of beads to a couple, even just one should be ok for widefield. Starting from Huygens version 2.16 it is less a problem if the PSF is somewhat truncated because of the build-in PSF extrapolation. This is also true for confocal PSFs which tend to be truncated always.

Am I right in assuming, that because microspheres are spherical, a mismatch between the refractive index of the bead and the surrounding medium is of no significance?

What happens when you use a small bead with slightly higher r.i. than the surrounding medium to measure the PSF that it will be not quite homogeneously excited. I think this effect on beads with a size approximately equal to the size of the diffraction spot in XY and so much smaller than the diffraction spot in Z can be neglected. The emitted light from within the sphere might seem to come from outside the sphere because of the sphere acting as a lens, but this is at most something in the order of diameter*( RI_bead/RI_lens -1) outside the sphere, so something like 10nm, also neglectable.

This is what you can do:

1. Close images you are not immediately working on and turn off the Undo system.
2. Use the crop tool to crop the data as much as possible, especially in the Z direction.
3. Use either QMLE or CMLE to deconvolve the image. If possible both will split the data into bricks and generate PSFs matching the bricks. In this way the computation of a single huge PSF is avoided. Currently, widefield images can only be processed brick wise if the data is sufficiently shallow compared to the NA. This is the case when the base of the aperture cone as truncated by the upper and lower planes is far smaller than the lateral extent of the data. This is often true, but overestimated NAs can spoil it. It helps to cut off as many z-planes as possible since this not only reduces the data size, but also allows more efficient brick cutting.
4. Lastly, make sure your system has sufficient swap space.

Dr. Markus Duerrenberger designed a confocal calibration kit, now available from Polysciences. This kit contains three different sizes of latex beads 0.2mu, 0.5mu and 1mu (all +/- 0.001mu), fluorescent in the whole range of wavelength from 280nm up to 647nm. These beads work extremely well for the Huygens Pro, but also all kinds of alignments and adjustments of the microscope can be done in a highly precise way.
Also Invitrogen and Thermo Scientific have good beads available. See also Where can I find a listing of available beads? to find a list of available beads.

The gensphere command generates a so called "bandlimited sphere", i.e. a geometrical sphere with no spatial frequencies above half of the sampling rate that you intent to use. If a perfect sphere should be used an unlimited number of spatial frequencies is involved and aliasing artifacts are generated due to Nyquist sampling violation. The ringing is a result of the sphere being bandlimited, i.e. perfectly antialiased. Removing the rings would mean corrupting the spatial frequency content which in turn would lead to a sub-optimal measured PSF. Because it is later convolved with a similarly bandlimited but smoothly rolling off PSF it is doesn't matter.

In typical conditions (1.3 lens, 488 excitation) good sampling is around 50nm x 150nm. Laterally 70nm is fine, you shouldn't go beyond 100nm, axially it is best to stay at 150-200nm. If you sample around the 70nm x 150nm you will see that the restoration is capable to reduce noise considerably. As a result you might need less signal than you thought before. If practical considerations (bleaching, data size) don't allow these sampling densities, you'll just have to do the best you can. In our experience, unless you undersample dramatically, the restoration will always improve your image.

See UnderSampling.

The aquisition zoom factor only affects the scanning parameters and as far as we know has no effect on the computation of the backprojected pinhole size. <br>
If the pinhole is specified in terms of Airy units the backprojected radius is computed as (from the FAQ):
backprojected_RADIUS = number_of_Airy_disks * 0.61 *
To compute the backprojected radius from the physical pinhole size see the tables in the Huygens Essential or Professional User Guide.

Oversampling is completely harmless. The more samples the better, though it could be argued that oversampling is not *necessary*. Of course there are practical reasons to limit sampling: object size, memory requirements, bleaching and so on.

If you have loaded an image in Huygens Pro, select it and select Edit-> Nyquist you get a report of the sampling situation taking into account all known parameters. Rule of thumb is for WF: 100nm lateral or better; around 200nm axially. 250nm would be fine too, but we distrust some z-motors with round numbers.

An undersampled image stack is a stack in which the Z or XY samping intervals are too large. Undersampling means that the sampling interval is too large to capture all information about the object generated by the microscope. In Huygens Essential the sampling values will be colored orange in case of undersampling. In case of severe undersampling the color will be red.

In Huygens Pro the optimal sampling density for the optical conditions under which an image is recorded can be computed by selecting its thumbnail and then Edit->Nyquist. When an image is recorded at this so called Nyquist frequency the digitized sequence contains *all* information carried in the signal. This makes it possible to reconstruct the image at any location, so not limited to the sampling positions. The Nyquist frequency is twice the highest spatial frequency (bandlimit) transmitted by the microscope.

There seems to be a discrepancy between the way Huygens defines stepsize and the way my microscope defines stepsize.

In the Huygens the stepsize (in any direction) is just the distance between samples, 'sampling distance'. Also, in all directions the numbering starts with 0, so the index for N samples runs from 0..N-1. This is the conventional way of indexing samples. Some microscope manufacturers use a special 1..N numbering scheme in the z-direction, but still 0..N-1 in the XY plane.

Yes, read our undersampling Is undersampling in Z-direction a serious problem?.

For a general discussion on correct sampling and aliasing see:

• Gonzalez, R.C. and R.E. Woods. (1992) Digital Image Processing. Addison-Wesley. ISBN 0-201-50803-6. p111 e.v.

For a discussion of microscopical bandpass characteristics see the papers below. Be sure to read both!

• Sheppard, C.J.R.The spatial frequency cut-off in three-dimensional imaging. (1986a). Optik 72 No. 4 131-133.
• Sheppard, C.J.R.The spatial frequency cut-off in three-dimensional imaging II. (1986b). Optik 74 No. 3, pp. 128-129.

For a practical discussion see also SVI's 'Deconvolution Recipes' manual or the Nyquist Calculator and Nyquist Rate.

No, it is a function of the optical properties of your system. It all revolves around the Shannon theorem, that states that for a bandlimited system (all our microscopes) it is totally sufficient to sample at the Nyquist rate. Now suppose the maximum spatial frequency passing a microscope is one cycle per 100nm (1.3 NA oil, confocal, 488/520nm, sampling at 50nm to sample peaks and valleys of the 100nm periodic wave). If you have a periodic structure of lumps spaced 80nm apart then this structure is not imaged, apart from its average value, nothing of it. Can't restore it, no way. If there is omly one single object and you know it is a sphere then restoration could consist of determining its center of mass. The accuracy of that depends on the SNR, but you could easily reach 10nm. Job done! The regular restoration procedure could also do it for you, but obviously to get such an accuracy in determining the peak location of the object you would have to resample the data to a higher sampling interval of 10nm. (You could also play it a bit dirty by not deconvolving with the PSF, but with the known image of the object; out comes a single peak where the center of the object is).

A more interesting object is for instance a two-blob object with a spacing at the Nyquist rate. Now the most interesting parts of the object spectrum are cut off by the microscope. The problem now is that the transmitted piece is the same for a whole family of objects. The family which has a spectrum quite like it is even larger. The restoration algorithm must now choose among them, the first selection being to exclude all objects with negative values. The better the SNR, the better the restoration algorithm can exclude objects of which the spectrum is slightly dissimilar to the measured spectrum. For confocals the situation is worse because they already attenuate everything beyond say 60% of the band practically to zero (depending on the pinhole). So in practice there is little hope for resolving objects at the edge of the band.

See Nyquist Rate and Nyquist Calculator. The following formulas can be used to compute the Nquist rate.
Widefield microscope:
Nyquist_lateral = lambda / ( 4 n sin(alpha))
Nyquist_axial = lambda / ( 4 n (1 - cos(alpha)))
with the wavelength, the refractive index of the medium (1.515 for immersion oil). is the half-aperture angle obtained with:
with the Numerical aperture. Many calculators use the 'sin^-1' or 'asin' symbol for the arcsin function.

Confocal microscope:
Assuming the excitation and emission wavelength are equal a confocal microscope doubles the bandwidth so halves the Nyquist sampling density.

Both Huygens Pro and Essential take the exact wavelength into account when computing the Nyquist rate. In case of multi photon excitation they also take the number of excitation photons into account. Both will color the background of X, Y, Z sampling density entry fields orange (moderate undersampling) or red (serious undersampling) when detecting undersampling. In Huygens Pro you can look up the Nyquist rate for a particular image by selecting it and Edit->Nyquist rate.

The Z-sampling is modfied by foreshortening due to differences in refractive index of the media and the immersion oil: the 'reverse fishtank' effect. In most cases the z-sampling as specified in the raw datafile is the nominal sampling distance, i.e. the distance the table or objective actually moved in Z without taking foreshortening due to refractive index mismatch into account. The mismatch induces spherical abberation and can have a profound effect on the PSF shape and effective aperture. The PSF generator takes all this into account.

After deconvolution the remaining geometric distortion can be corrected by multiplying the z-sampling distance by the ratio of the medium and immersion refractive index, a number in most of the cases < 1. See also Fishtank Effect

To find out the ideal sampling, you can always use the online Nyquist Calculator, entering the image parameters (including the number of photons: 2). You can also use the Huygens Software in your computer. To find out how large are your samples in relation with the ideal Nyquist Rate, do the following:

In Essential

• If you have an image for which you want to compute the Nyquist rate, open it and check its parameters (right click -> "Show parameters").
• Whenever you change the Microscopic Parameters of an image, the Nyquist rate is recalculated. Modify the image parameters to match the microscopic conditions for which you want to compute the Nyquist rate, then check the parameters again (right click -> "Show parameters").
  

In Professional

• Select the image, select Edit -> Nyquist rate in the main menu. You'll get a popup displaying the Nyquist rate.
  

See also Multi Photon Microscope.

If in 2-photon excitation a pinhole is not used it is the excitation distribution which determines the imaging properties of the microscope and therefore the Nyquist rate. The excitation intensity field is that of a widefield microscope, but since due to the 2-photon effect the effective excitation distribution is the square of the intensity distribution, the imaging properties are vastly different. The squaring operation makes the distribution more 'peaked', resulting in an improved resolution. It also causes the bandwidth (and with that the Nyquist rate) in x, y, z to be twice that of a widefield microscope at the same wavelength.

Importantly, the 3D shape of the band-pass area is very different: while the widefield area has a wedge at the center causing the large widefield blur cones, the 2-photon bandpass area has no such defects.

See Pinhole And Bandwidth.

Old information
In principle, the Nyquist rate is independent of the pinhole size. This is due to the choice to relate the Nyquist rate to the theoretical bandwidth of the system: the spatial frequency beyond which *nothing* is transferred by the microscope. It would be a different story if we would have used a criterion based on attenuation of spatial frequencies below a certain factor. (Larger pinhole sizes attenuate higher frequencies more, but still are not zero). Although a practical approach (because about last third of the band has so low intensities that they can be considered zero most of the times) this involves an arbitrary choice, so therefore we base the Nyquist rate on the well defined theoretical bandwidth.

A problem occurs with extremely large pinholes like those used in two-photon systems. In these cases the optical properties are practically identical to a widefield system whereas due to the presence of the pinhole the theoretical bandlimit is still that of a confocal microscope. In the two-photon case it is best to set the microscope type to 'widefield' when doing deconvolution with the Huygens Software, since this will result in the same optical properties but with a more practical Nyquist rate. Note however that with single photon confocals even a very large pinhole will still have a noticeable effect on the blur contributions of far off focus regions, thus improving resolution along the optical axis.

The Confocal Microscope provides more information than the widefield system; in theory the confocal Sampling Density should be twice the widefield density. This holds for in all 3 dimensions, so you'd get 8 times more voxels.

A 'typical' widefield setup (1.3 effective NA lens, 500nm emission) is sampled well at 100 nm laterally, 300 nm axially. However, due to good SNR ratio's deconvolution can often gain a lot in Z, so you might as well go for 200 nm. With a 100x lens and a CCD with 6.7 micron cells you get 67 nm laterally. (This is assuming that there is no extra magnification; otherwise the total magnification must be used when calculating the pixel size). If this bloats your data too much you can try binning to increase the lateral size to 134 nm, but you will already start seeing some 'staircasing' effects on thin filaments in the deconvolved image (Aliasing Artifacts).

In the same typical confocal case a nice sampling rate would be 50 nm in Z, 150 nm axially. In case of bleaching problems you can stretch this up to 75 nm lateral, and after that increase the Z-sampling.

If the Parameter Editor or the Parameter wizard starts coloring the sampling fields orange, then you start to undersample; red means severely undersampled. See Nyquist Rate.

This greatly depends on the optical paramaters, in particular on the NA and microscope type. Together with the wavelenghts and refractive indexes these determine the so called Nyquist sample distance, the maximum sampling distance at which all image information is captured. For a plot of the dependancy of the Nyquist rate on the NA and microscope type see the User guides, or go to Nyquist Rate. To calculate this figures online see the Nyquist Calculator.

• Confocal microscopes: While sampling at the Nyquist rate is a very good idea, it is in many practical situations hard to attain. In these cases larger sampling distances may be used. For confocal images with a Airy disk sized pinhole the lateral sampling distance may be up to about 1.6 of the Nyquist distance. When much larger pinholes are used, up to 2x.

When very small pinholes are used to optimize resolution, or any other case where high resolution is required, we do not not recommend undersampling since this would defeat the purpose, and since it might limit the deconvolution result.

• Two photon microscopes: Two photon microscopes without a detection pinhole (e.g. non descanned systems) follow the rules of a small pinhole confocal microscope relative to the longer excitation wavelength. The addition of a detection pinhole increases the theoretical bandwidth, but since usually large pinholes are used the practical bandwidth increase is small.
• Spinning (Nipkow) disk microscopes: Regarding sampling criteria spinning disk microscopes behave like confocal microscopes with, depending on the disk, a fairly large pinhole. For a typical system we do not recommend more than 1.6 times undersampling.
• Widefield: Widefield data are more sensitive to undersampling; stay below 1.3. In case of low numerical apertures like 0.4 we recommend not to undersample in the axial direction.
• STED : The remarkable property of STED microscopes is that they do not have a band limit in the strict sense. That means that any sampling rate is a compromise between practical considerations and the to be reached resolution. Since lateral STED HIW resolution can achieve 50nm in good conditions we recommend sampling around 25nm. In difficult condition this can be increased, but that will limit deconvolution.
• Multi channel data: The sampling rate should be derived from the highest resolution channel.

The actual and the ideal sampling distances of an image can be seen in Huygens Essential right-clicking on the image thumbnail and choosing 'Show parameters'.

To compute the ideal Nyquist rate of an image in Huygens Professional, select the image and choose Edit -> Nyquist rate.

Widefield and confocal microscopes differ in the amount of information they are able to extract from a specimen. One way to express this is to look at the finest details or highest spatial frequency the microscopes transmits: for a confocal microscope this is nearly twice as much as in an equivalent widefield microscope, in all directions.

According to the Nyquist theorem a signal should be sampled at twice its highest bandwidth so confocal microscopes need twice the sampling density of widefield microscopes. Although the confocal microscope is able to transmit twice as fine details as the widefield microscope, it attenuates these very strongly. Beyond say 60% of the highest frequency practically nothing is transmitted, especially for not-ideal pinhole cases. Therefore, while sampling according to Nyquist rate remains the safest solution, in the case of confocal imaging it is defensible to reduce the sampling rate to about 60% of the theoretical rate, for example in typical condition one sample per 50/0.6 = 80nm.

In the widefield case, high spatial frequencies are also attenuated as the band limit is approached, but to a much lesser degree than in the confocal case. Therefore we do not recommend to stay below 1.3 of the Nyquist rate. In case of low numerical apertures like 0.4 we recommend not to undersample widefield images in the axial direction.

A practical example:
Assuming a 1.3 N.A. objective lens and 488nm excitation, 520nm emission you need to sample around 50 x 50 x 165 nm to get an optimally sampled confocal 3D image; 100 x 100 x 330 nm in the 3D widefield case. As mentioned above, widefield images are more sensitive to undersampling, i.e. a violation of the sampling rule has a more dramatic effect on widefield images than on confocal images.

For more details on sampling densities for other microscope type, see the FAQ What is the maximal voxel size at which Huygens can still do a good job?.You may also want to have a look at Sampling Density and Nyquist Rate.

If your CCD camera has the option of binning, the binning can be used to increase the signal at the cost of resolution. With binning the intensity of several pixels is integrated to form a super pixel. For instance, 4 x 4 pixels are summed to form a bigger and brighter signal at the cost of resolution. When binning is applied the effective voxel size increases.
Example: If the actual detector element size is 6 micron and a 2 x 2 binning function is active, then the effective pixel size is 12 x 12 micron. As a consequence the magnification has to be increased by a factor of 2 relative to the non-binning mode in order to satisfy the sampling criteria.

Indeed, lowering the NA to allow for larger samples is the correct way. The drawback is that the axial resolution of the image will decrease with the square of the reduction of the NA. The sampling density as function of the NA can be found in the Huygens User Guides in the chapter Establishing Image Parameters.

See Biorad MRC_500_600_1024 and Biorad_Radiance. Old information: from communication with Brad Amos we learned that the factor for the 1024 is 53 and for the Radiance is 60. We had feed back from a customer who reported "Using Brad Amos' value, the deconvolution is working well".

For the most used Yokokawa disk spacing is 250 micron, so with an 100x lens backprojected about 2.5 micron This you can check by stopping the disk.
The pinhole diameter is probably 25-50 micron, resulting in a backprojected radius of .125 - .25. The latter is about an Airy disk.

The Perkin Elmer Ultra View is a Yokagawa disc and the Yokagawa disc is always a Nipkow disc. You must choose the Huygens option for Nipkow disc microscopes.

A system which refuses to pass spatial frequencies beyond a certain limit: all optical instruments. See Band Width.

This question can be particularly relevant for widefield microscopy. Widefield images are fuzzy in the neighborhood of the in-focus region, which rises the question of how far above and below the in-focus region (Z direction) the image should be recorded to ensure an optimal deconvolution result.

For a good widefield deconvolution it is not necessary to acquire large regions above and below the in-focus zone. In widefield microscopy the blur can spread through large regions. The Huygens algorithms take this into account, thus imaging very far above and below the in-focus region is not necessary to achieve good deconvolution results. The image can be cropped far from the in-focus region to speed up the deconvolution process. To avoid cropping slices that contain useful information, leave a border of around 1-2 Half Intensity Width sizes. As an example, in widefield, well-sampled images recorded with NA 1.3 this corresponds to 0.75 to 1.5 microns.

A confocal image deconvolution already benefits from including a few planes. Even including one neighboring plane on each side of the plane of interest gives already an improvement in resolution and signal. However, if a realistic representation of an object in the plane of interest is required, it is advised to image at least a Z-stack that matches the dimensions of the PSF. You can calculate this with the calculator on NyquistCalculator. For larger objects you would want to have at least a number of Z planes that matches half the axial width of the PSF on both sides of the object.

As a rule piezo-electric focusing devices have far greater positioning accuracy. For example, older stepper motor driven stages already have trouble positioning with a 0.5 micron accuracy while good piezo stages are better than 10nm. On the other hand, modern stepper motor setups claim 0.1 micron 'accuracy' which in principle is sufficient for many applications. Whether the accuracy is really sufficient depends on factors like the maintainance state of the microscope, age of the lubricants in the gears, and so on.

Piezo driven focussing systems are also not without problems: hysteresis, temperature drift to name a few. To compensate for the latter, especially in cases where experiments are to be conducted at 37 degrees, we recommend a sensor feedback system.

If you are using a monochrome camera you have one channel to which you assign the peak of the fluorescence emission spectrum. If that peak is cut off by the bandfilter, the effective emission wavelength at the detector will be close to the filter cutoff wavelength.

If on the other hand you are using an RGB camera, the situation is a bit different. Each of the camera components has its own, probably not so sharp band filter. The camera filters are in cascade with the microscope bandfilter. Suppose the microscope filter passes all wavelengths longer than 500nm, and the blue camera filter passes all shorter than 500nm, then the Blue channel will be dark. In a different case, suppose the microscope filter passes all wavelength longer than 570nm, the green camera filter from 500-600nm, and the red camera filter from 580nm and longer. Then both Green and Red channel contain an image. With little loss of accuracy these could be added and deconvolved at 570nm emission wavelength (you need Huygens Pro for that).

In the most likely case two of the three R, G, B channels will contain only a very dim image. You might as well discard these and concentrate on the brightest image.

The embedding medium in which the sample is mounted. We also include an immersion medium field to account for water immersion lenses.

Start with the smallest pinhole. This gives the smallest PSF resulting in the most conservative restoration (less artifacts). If you are not satisfied with the resolution you may wish to increase the pinhole size.

As a rule, it is never a good idea to tinker with the greyscale of an image prior to deconvolution. For example, if one would clip the bright pixels in an image the in-focus image of the object will not correspond anymore with the non-clipped blur it causes. In the restoration this usually results in insufficient blur removal.
A different thing applies for Hot Pixels. These can be removed effectively with the Hot Pixel Remover.

Arcsin, a.k.a. asin or sin^-1, is the 'reverse' of the sin function:
if then for .

For more information see BackprojectedPinholeCalculator.

The formula is:

with r the backprojected radius in nm, d the pinhole diamater in mm, the system magnification, the objective magnification. The "2" is for diameter to radius conversion.
Depending on the configuration the system Magnification for the MRC 600 or 1024 is 53-83. The system magnification of the Radiance is reported to be 60.
For more information and a calculator you can visit BackprojectedPinholeCalculator

The message means that a refractive index of 1.14 can't be combined with an NA of 1.2 because according to Snell's law NA = r.i. x sin(alpha) and sin(alpha) <= 1 so NA < 1.14 when r.i. = 1.14 for any lens.

If you specify an NA which is higher than the r.i. of the medium, then this implies total internal reflection at the glass/medium interface, causing the oblique angles of the excitation light to be reflected back into the microscope, thereby reducing the effective NA.
See also TotalInternalReflection.

I intend to obtain some diatoms which have detail in the sub-micron range and which have 3-d structures. It should be possible to image them by immersing them in a fluorescent medium and model their appearances using different algorithms to find which one gives the best approximation of the real appearance of the diatom (as determined by scanning EM).

Imaging diatoms is indeed a difficult task. This is so because the skeleton of diatoms has a high refractive index and can seriously spoil light propagation, resulting in a PSF which varies strongly from place to place.

It is a good idea to do this. Often we see considerable shape dependencies in confocal PSFs, also in widefield PSFs.

The detector is seeing a broad spectrum of light, exactly what this is will depend on the filter characteristics prior to the detector. We once did some simulations where we integrated the PSF(s) over the wavelength band of the dye spectrum and compared that to the PSF of an 'effective' wavelength: a last decimal case. A good estimate of the effective wavelength is the centre of mass of the spectrum, but since the main part of the spectrum is roughly symmetrical and effect of a slight error on the PSF shape is minimal, the peak value is also fine, or the mean between bandwidths. In case of long pass emission filter, there is no mean (one bandwidth goes to infinit), and the peak may not be the correct. In that case the center of mass needs to be estimated. Read more at Emission Wavelength where images explain the difference.

The laser line: that is the light actually going through the optics.

Refractive index for vectashield: 1.457

In the form for optical parameters, there are two fields for Refractive Index. One is for the oil (oil immersion objective), the other for the medium. Is this the medium in the samples like DAKO or glycerol? The other, the medium between the samples and objective?

Yes! If you enter a figure not much lower than the NA you'll see it gets highlighted to warn you against an optically poor situation.

Low light level images of say maximal 25 photons per pixel will naturally yield pixel values which are low with respect to a ccommon range of 0.255. That does not mean the image is poor in the sense of a poor Signal to Noise Ratio (SNR). Since SNR = sqrt(maxPixelCount), it is easy to compute the SNR from a photon counted image: in this case 5. Images with SNR 5 can usually be quite well restored.

A measured 1 micron latex bead was sampled at 100nm in all directions. In the original image (assuming typical microscope conditions, 1.3NA, 500nm excitation wavelength) the bead should have an axial size of roughly 1.8 microns axially and 1 micron laterally, so 18 samples axially and 10 samples laterally.
A factor which can increase this elongation considerably is refractive index mismatch of embedding medium and coverslip. If this is the case and you can't increase the refractive index of the medium you might consider purchasing a water or glycerin immersion objective. To get a more accurate estimate of what the bead should look like in the microscope, you can simulate the imaging process with Huygens Professional:

• start Huygens Pro
• generate a sphere + do an FFT on the result
• generate a PSF + do an FFT on the result
• multiply the two and put the result in say image 'c'
• do a reverse FFT on 'c' (with the FFT tool in auto mode this will go automatically)
• do a 'to/from optical representation' (restoration menu) to shift the origin of the image to the center.
• to make things realistic you can throw in some Poisson noise.

The result should not be too different from the measured image.

In order to generate or reconstruct point-spread functions Huygens needs to know the excitation wavelength in nanometer. For multiple-photon excitation use the actual wavelength (e.g. 780 nm) and specify 2 for the "number of excitation photons" in the parameter dialog. The model assumes that all participating photons have the same wavelength.

I am using Leica TCS SP2 confocal microscopy. The pinhole setting is 1 Airy, which is about 82um. This information is obtained directly from Leica software. But When I was using Huygens essential, it told me the pinhole size is too large. I also looked at the users guide, found this formula r=0.61*lambda*N/NA. According to it, I got 238nm. Could you tell me what I should choose?

The pinhole measure as used by Huygens is the "backprojected pinhole RADIUS", which means that the physical size should by divided by the systems magnification factor. Using a 100x objective lens the value is 410 nm, which is still pretty high (250 nm is a common value), so maybe there is some more internal magnification in the system we are not aware of.

Using the Airy-disk unit prevents you from having to know about magnification settings so 238 nm is the one to use.

Some confocal microscopes report the pinhole size with the Airy disk as unit. Compute the backprojected pinhole radius from the following formula:
backprojected_RADIUS = number_of_Airy_disks * 0.61 *
NA = the numerical aperture, number_of_Airy_disks = the number of disk diameters fitting in the pinhole diameter.
The Airy disk diameter is the diameter of the first dark ring of the diffraction pattern in the focus of a lens, the so-called Airy disk.

When light passes from a dense medium (glass) into a less dense medium (water) total internal reflection occurs above a certain angle of incidence. High aperture objectives easily generate rays at angles beyond this limiting angle. When the refractive index of the medium is below the objective's NA this will occur.

Example: with an NA = 1.4 and a watery medium (refr. index slightly above 1.3), the most oblique rays cannot pass into the medium and will be totally reflected, effectively limiting the NA of the objective.

See also: What does the message: 'total reflection at glass/medium interface (...)' mean?
And see the wiki article Numerical Aperture.

Find out the pixel sizes of the camera's CCD chip. Typical values are 6.7 x 6.7 micron. With a 25X objective the XY sample sizes will be 6700/25 = 268 nanometer.
This is assuming that there is no extra magnification. If the microscope has internal magnification that should must be considered. In that case, the total magnification (all magnifications multiplied) must be used when calculating the pixel size.

Customers noticed that small microspheres undergo Brownian motion in glycerol and water. We have heard of people using polylisine to attach the beads to the glass, but there are probably a dozen more tricks we are not aware of.

Very large or no pinhole

If there is a non-descanned detector (NDD), or a very large (say > 10 Airy disks) pinhole, the contribution of the pinhole to the image formation can be ignored. In this case the microscope type can be widefield in combination with 2p photon. This will result in more realistic values for the Nyquist rate than a confocal setting.

Fairly large pinhole

If there is a pinhole in the order of a couple of Airy disks, typically 500nm backprojected radius, it must be taken into account. So in this case the microscope type should be set to confocal.

We have seen this many times. Please see Banana Psf.

It is more like estimating. Just look at the angle of the blur-cone fanning out from the bead along the optical axis (Z). The largest angle is the aperture, whose half (α) you need to use in the Numerical Aperture definition. Usually it can be easily seen in XZ slicing mode. A 1.3 NA oil lens should show α = 60 degrees with the optical axis, making the total cone angle 120 degrees. Often it is less!

It is a sure way to spot optical or calibration problems. The shortcoming of this method is that you can't discriminate directly between errors in the sampling calibration or the effective NA. However, if there is remarkable asymmetry then it could be explained by Spherical Aberration (see Mismatch Distorts Psf). Assuming the lens is good, this must be caused by a Refractive Index Mismatch. In turn that will lead to a reduction in effective NA. So in these cases a refractive index mismatch is at least one of the factors causing the problem.

There are many different types of agarose and gelatin, and they may not be equally suited for embedding of light microscopy samples because of autofluorescence, light scattering etc. Are there any special brands of material or protocols/publications for purification/embedding available? Or any other material keeping suspended samples (like beads) in place in three dimensions?

I use gelatin to simply immobilize my sample (suspend it in place in 3-dimensions) which will otherwise tend to "swim away" in an aqueous medium. I do not notice any autofluorescence in the 5% gelatin.
(Answer by Harry Leung, M.Sc.,Technical Officer, Zoology Department, University of Western Ontario, London, Ontario N6A 5B7, E-mail: leung at julian.uwo.ca)

Please read the page which explains the module options.

The axial resolution of a confocal microscope is inversely proportional to the pinhole size. However, as the pinhole becomes very small, so does the detection efficiency. Generally a good compromise between sensitivity and resolution is found at a diameter of one Airy disk unit.

In practice, depending on wavelength, for a 1.3 NA objective this corresponds to a backprojected pinhole radius of 200nm to 300nm.

The SNR describes the ratio between signal and noise in the image. The deconvolution algorithms require the SNR of the image as an input parameter. If the deconvolution algorithms are provided with an SNR value that describes the image accurately the restoration will be optimal. However, if the provided SNR value is too high, the image noise will be amplified. If the SNR value is set too low, the noise will be reduced at the cost of the final resolution of the image. Therefore, the SNR can also be thought of as an artifact limiter as far as the deconvolution is concerned.

Confocal images

In many confocal images the SNR is due to photon noise. If the background is mostly zero with spikes here and there, then these spikes are probably single photon events. From the height of the average small spike one can roughly estimate how many gray levels correspond with one photon. If, for example, such a single photon event has an intensity value of 5, then the maximum intensity of the image (for example 255) corresponds with 51 photons. The SNR around the maximum intensity area will be: square_root(51) = 7.

With a good confocal image, and when using an 8-bit converter, one can easily get into a situation where one gray level corresponds with more than one photon. In such case, the above procedure fails, but one could still start a run with SNR 20-30 and increase it later on.

Some confocal microscopes are equipped with photon counters like avalanche photo diodes (APD). In this case the SNR is simply the square root of the brightest part of the image.

Widefield images

Images from widefield microscopes equipped with 12-bit CCD cameras usually have an SNR in the range of 40-60. These kinds of images can also be deconvolved with the fast Quick-MLE algorithm for low noise images.
The Huygens Remote Manager is equipped with an automatic SNR estimator free to use here (registration needed). See also Set The Signal To Noise Ratio.

Experimenting with the SNR

In practice, the SNR can be estimated out by calculating the number of photons in the image, or by rule of thumb reasoning. One can then start a first deconvolution with that SNR value and inspect the result for artifacts. If no artefacts are visible, the deconvolution can be continued with a higher SNR setting (say 5-10 higher). If the results are good but there is still a residue background, the deconvolution can be continued with the same SNR value but with a higher background setting.

Noise generators are used to generate realistic synthetic images to be used in Imaging Simulation and test runs. Moreover, the addition of noise can be used to visually mask areas of an image where the data has overflowed (bright patches).

See pnoise and gnoise.

Yes, indeed. The SNR being the square root of the maximum photon count would, in this case, yield: SNR = SQRT(256) = 16.

As a very basic approach, the following values can be used as a first estimation of the SNR of different image types:

• Bad quality confocal image: SNR = 10
• Noisy confocal image: SNR = 20
• Good quality confocal image: SNR = 30-60
• Good quality widefield image: SNR > 40

If the image noise is clearly amplified after the restoration then the SNR value provided to the deconvolution algorithm was too high. On the contrary, if the restored image looks too smooth then the SNR value was too low. In both cases, please proceed to restore the raw data again with an adapted SNR value.

See Set The Signal To Noise Ratio.

Too high SNR setting in both MLE and ICTM methods may lead to artifact generation. These artifacts have a different character in the MLE and ICTM method, so they will be treated separately.

• Artifacts in the ICTM method due to overestimation of the SNR: There are two sources of artifacts with this class of method: amplification of noise and 'ringing' artifacts, i.e. fringes around sharp edges. Ringing artifacts are greatly suppressed in the ICTM procedure. Moreover, when you switch on 'ringing reduction' the ICTM procedure will lower the SNR setting when it detects the onset of ringing. The drawback of this feature is that it increases the data dependency of the procedure. So, when you are planning to restore and compare a number of images of the same type, we do advise you not to use this switch. More troublesome is noise amplification. The amplification increases with the number of iterations. Most notably it shows up as semi-periodic structures in the background. The ICTM method can be used with the background correction option. This enables you to set as background value the average of the image background. Use Analysis->background from the Operations window to inspect the background. This feature will enable you to remove the background including the ringing phenomena effectively. Conclusion: overestimation of the SNR (say by a factor of 2) won't do much harm. However, since noise amplification artifacts may also occur inside the object, be careful with high SNR values (>100) in combination with large number of iterations (>100).
• Artifacts in the MLE method due to overestimation of the SNR: Due to the built-in background correction and its optimal treatment of background (nearly always Poisson type) noise the MLE method is hardly sensitive to background noise amplification. For noisy images, amplification of noise within the object is also less than in the ICTM method. Overestimation of the SNR value in noisy images will result in reduced noise suppression. A potential danger of overestimation of the SNR parameter (>100) in low noise images is 'over-restoration', i.e. objects are estimated too small. This occurs in combination with high iteration counts, >100. With settings of the SNR parameter below 30 there is not much danger in over-restoring the object. Conclusion: for restoration of high noise images the MLE method is quite safe: there is a straightforward trade-off between noise and resolution gain.

In both the Maximum Likelihood Estimation ( MLE) and Iterative Constrained Tikhonov-Miller (ICTM) methods this will lead to loss of detail in the result. The ICTM method with SNR < 10 and MLE with SNR < 5 will actually lower the lateral resolution. Axial resolution will be reduced in the ICTM method with SNR < 10; in the MLE method with SNR < 3. The figures given here are estimates, they depend on the data. See Set The Signal To Noise Ratio.

You can install a license by using any Huygens Product and going to the Help -> License menu to add, delete, explain and check the license. Start the software by clicking its icon or typing the corresponding command at the Unix shell prompt. All products use the same license file, thus it does not matter which product you use to install a license string.

On Unix systems, you can also add the string with a regular text editor in /usr/local/svi/huygensLicense. On Mac OS X it might be in a different place, depending on where you installed the software. Typically that would be /Applications/SVI/huygensLicense. You probably need to have root access to edit this file. On a Windows PC it is by default 'Program Files\SVI\huygensLicense'. Best put the new license above the old ones.

On Unix systems you can also store the license in a text file named huygensLicense in your home directory. This is especially useful on Mac OS X.

They can be shared. The license reader stops after the first valid license for a particular date and machine id. It is not necessary to share /usr/local/svi: if the SVIHOME environment variable is set it follows that.

You can download the latest releases and patches available on our download page http://www.svi.nl. The existing license will work as long as the license product number is the same as the number of the Huygens release:

e.g.
HuEss-4.2-wcnp-d-ftv-emnps-eom2012Oct01-f2372e2f612c8cf3-{your@email.com}-ab80d315d5bf4aa89d2c

or

HuPro-4.2-wcnp-d-ftv-emnps-eom2012Oct01-f2372e2f612c8cf3-{your@email.com}-ab80d315d5bf4aa89d2c

or

HuScript-4.2-wcnp-d-ftv-emnps-eom2012Oct01-f2372e2f612c8cf3-{your@email.com}-ab80d315d5bf4aa89d2c

The above licenses are valid for the Huygens 4.2.XpX versions. Huygens version 4.3.XpX or higher will not work with 4.2 licenses or lower. When there's a new release customers with valid maintenance receive new licenses updated for the new Huygens version.

Yes, for Mac and Linux you can specify a different installation location than the default '/' with the '-r' option to the command line installer 'inst' or the software manager 'swmgr'. Example from your home directory:

inst -f dist65 -r TestInstall

(this assumes the distribution is in 'dist65'). This places the executable in

~/TestInstall/usr/sbin/huygens2

In addition you need to set the SVIHOME environment variable, in this case to

~/TestInstall/usr/local/svi

The license string should be in '~/TestInstall/usr/local/svi/huygensLicense'.
To run huygens type

~/TestInstall/usr/sbin/huygens2

When the Installer program gives the error: "The install package cannot be opened. The Bill of Materials for this package was not found." it tells us that certain needed files inside the installer cannot be found.

It seems that this error can have more than one cause. Probably the most common reason is a corrupted installer file. The file could have been corrupted during transfering from one computer to another.

The following command sets the number of cpus:

huOpt cpu ?-min integer? ?-max integer? ?-mode <compliant|ignore (compliant)>?

By default the system complies to values suggested by the system. If the mode is set to ignore Huygens will try to force the requested number of threads from the system. E.g. to limit the number of cpu to 4, run the following in the tcl-shell:

huOpt cpu -max 4

The Huygens compute engine (CE) uses the OpenMP standard for multiprocessing. The multiprocessing behavior of the CE can controlled with OpenMP environment variables or from the Huygens-Tcl layer.

Environment variables:

• The number of threads to be used is controlled through OMP_NUM_THREADS, for example: 'setenv OMP_NUM_THREADS 2'
• Allow the operating system to dynamically adjust the number of threads depending on system load: OMP_DYNAMIC (TRUE, FALSE), for example 'setenv OMP_DYNAMIC TRUE'. If OMP_NUM_THREADS is set then the number of threads varies between 1 and OMP_NUM_THREADS.

Huygens-Tcl layer:

huOpt cpu ?-min <integer>? ?-max <integer>? ?-mode <compliant|ignore>? ?-query <system|min|max|mode>

This allows setting of the minimum and maximum to the number of threads. Mode compliant is equal to OMP_DYNAMIC TRUE. With -mode ignore the CE tries to always use the max thread number. With the -query mode the curent state can be queried.
Example: huOpt cpu -max 4 -mode ignore to run on four threads regardless of system load.

Yes. All versions of Huygens are multithreaded.

If you view an image with the slicer, for each update a 2D image must be sent over the network. The amount of pixels in the 2D image is determined by the number of pixels in the slice, not the display size on the screen. The images are in full color (RGBA) which means 4 bytes/pixel. So, for a 512² image it is 1 Mbyte per plane, i.e. one second at best over 10Mbps ethernet. This is indeed roughly the update rate we get when running Huygens over a 10Mbps ethernet at SVI. So for smooth slicing with slice planes in the 1Mbyte range a 100Mbps connection is required.

To enable you to operate the Slicer over much slower networks the update rate is measured by the software. If the update rate falls below a certain threshold, currently about 0.5 s., the Slicer switches from smooth scrolling to jump scrolling. In the latter mode, the screen is only updated after you release the slider.

This means you have run out of swap space... You could of course increase the swap space by adding a swap file but it is also possible to have Huygens consume less memory by switching off the undo/redo system. Especially when running scripts it might not be really necessary to have an undo function. It is also good practice to destroy all images you have created in the script as soon as you don't need them anymore.

Some possible sources of error:

1. The string was not pasted in in its entirety.
2. The string got corrupted, for instance because it got split into two lines, see below.
3. There is a mismatch between the software release and your license.
4. You do not have sufficient rights to install the license, usually you need administrator rights, see below.
5. The system ID was for a different computer, see below.
6. The system's clock is not set to the correct date.

Typical problems when copy/pasting license strings are:

• The email client has split the string in different lines. The license string must be a single line, without any space or line break.
• The email client has converted a double dash (-+--+-) into a single long dash (—). A valid license string contains only 7 bit ASCII characters: basic English letters (a-Z or A-Z), numbers (0-9), dashes (-+-+-), at signs (@), curly braces (-+{ }+-), dots (-+.+-) and maybe other very basic symbols like underscores (-+_+-).

Normally you must have 'administrator or root rights' to install the license and will get an error message if you try to install the license without the proper rights. However, some Windows versions do not give a warning while afterwards the message cannot find a valid license keeps popping up. Right-click on the Huygens icon, and then choose 'run as administrator' and log in. Installing the license now will be successful.

Maybe the license string you are using is syntactically valid, but not intended for your computer. License strings are restricted to work only on a single computer, identified by its System Id. This ID is generated based on serial numbers of some of your hardware components. If you change any of these the System ID may change and the License Strings generated for it no longer work. A typical problem is caused by the change of a Network card. Consult your maintenance contract and Contact Svi to see how to update your license strings in such a case.

Does your license string not enable all the expected functionalities? The license string as used by SVI has the same appearance on all supported platforms. For each product you need to have a license string installed. Select an installed license string in the License manager and press the Explain license button. All details for the current license will be listed. If you run into licensing problems you may use this information to analyze the problem.

See wiki article → License String and License String Details.

A license entitlement represents a proof of purchase of a particular SVI software product. It entitles its owner to a single permanent, node locked license. It can only be used once. The license entitlement consists of an unlocked license string and a unique system ID. It can be requested here, or by sending the systemID together with name, organization and address information by email to license at svi.nl. Alternatively, this information can also be faxed to +31 35 683 79 71.

An example license entitlement for:

• Huygens Essential with
• all microscopic options (widefield, confocal, nipkow and two photon),
• a time option,
• issue date 2012 Dec 16,
• system ID of E123245

looks as follows:
HuEss-2.4-wcnp-d-t-emnps-2012Dec16-E12345-{license@svi.nl}-d852dfbe431e5bcb9317

You can download both RPM and Debian-style distribution packages from the download page. Simply double-click on the package or run the package installer. For debian, this is dpkg -i, for rpm, this is rpm -i install.

More information can be found at RemoteDisplay

More information can be found at RemoteDisplay

To view a surface rendered cut of an image, you need to cut off a piece of the image. This can be done with the Object Analyzer, where you can cut off any piece out of your image.

• You can create an ROI of the region you want to remove. Either by selecting a region with the selection tool or by using objects and anchors from the menu ROI -> Set.
• When an ROI is created, go to ROI -> Dataset cropping -> Remove intensties inside ROI and then export the image using the first button in the Data section at the left. Choose any setting, e.g. Overwrite All
• Visualize the new image with the Surface Renderer.
• When you have chosen Append to original, you can do some nice transitions in the Movie Maker, by increasing the transparency of one channel and decreasing the other the cut-off will be slowly displayed. Just play with it and explore all its possibilities.

If you do not have the Object Analyzer, you can use the cp command to copy a part of an image in Huygens Professional. Replicate the image with the repl command and clear the new image, such that it only contains zero values. Then copy any arbitrary block of your image to this new image with the cp command. For example, if you want the cut of a 100x100x100 image in the x-direction, with the left part deleted:

a cp -> b -dest {50 0 0 0 0} -src {50 0 0 0 0} -span {50 100 100 0 0}

See also: cp and repl

Yes, this can be done in Huygens Professional and Huygens Scripting. Suppose your image has intensity values from 5 to 100. You can stretch these values from 0 to 255 by using the cs command.

In the Operations window of Huygens Professional, go to Manipulation->Copy&Stretch. The Tcl command cs is used as follows:

a cs -> b -fromzero

When labelling an image, there are some limits to be taken into account. When these limits are reached, the software will give an error and report which limit is reached. These limits can be reached in case of a very noisy image, in which case filtering or deconvolution is necessary first.

There are two limits:

• The labels are saved in a 16 bits signed integer format. So 2¹⁵ - 1 = 32767 is the maximum number of labels (objects). That seems to be very much, but is not enough in the case of a very noisy image where a large amount of objects will be detected. Filtering or deconvolution is the solution here.
• The size of objects is limited to 10⁵ pixels. In case this limit is reached, try to use a higher threshold to make the object smaller.

See Set Channel Colors.

Yes. You can enlarge the Operations window and slide the sash (moveable border between the image on the left and the tabs on the right) to the right. The extra space is then filled up by the image, which gets larger.

Yes, there are basically three ways, in order of simplicity, depending on whether you want to use the gui or scripting.
Using a GUI
You can directly crop an image using the crop tool, which can be found in the buttonbar, and under Edit -> Crop. Or with the shortcut Alt-B. At startup the Intelligent Cropper estimates the region you will need for deconvolution. Read more at Intelligent Cropper. In Essential the cropper is found under Tools -> Crop...

Using scripting - crop symmetrically
In case you want to crop using the Tcl Shell or within a script you can do that by removing a symmetrical border. Use the add/remove border command, border. The size of the border to be removed can be specified as an option of this command, for instance '-100 -120 -40' to cut away 50 pixels in X on each side, 60 in Y and 20 in Z. Type <imagename> border ? to see how the command is used.

Using scripting - crop any part
In case you want an arbitrary part of an image you should first create a destination image which has the exact same size as that part. There are two methods to do this:

1. One way is to use the repl command to create a copy and use add/remove border to trim it down to the desired size. In this way the microscopic and other parameters of the destination are surely correct.
2. The other way is to create a new image with the desired size and then call for the adopt command to copy the microscopic parameters from the original to the cropped image.

After the new image is created, then use the cp command (see Operations windows under Arithmetic->Copy image part) on the original and specify source, destination and span as follows:

• source: the coordinates of the origin of the region you want to take out.
• destination: leave at 0 0 0 0 0(default)
• span: set to a size as large or larger than the sizes of the destination image. For instance if the destination has size {100 100 40} span could be {100 100 100 1 1}. The last '1' (ones) are for the (not used) time and channel dimensions.

See also: cp and repl

See the answer to the question: Is it possible to crop a stack with Huygens Professional?

Due to the band limited character of the image small negative values might show in the sphere. The spheres are stretched so the originally black "0-values" are stretched to some small non-zero value. When viewing the image in false colors, the background is not the lowest color and will thus appear in purple.

The Colocalization Analyzer is an interactive colocalization tool present in Huygens Essential and Huygens Professional, allowing you to obtain not only global image information, but also to quickly determine the properties of the different colocalization regions in your data.

CoLocalization analysis gives information about the amount of spatial overlap between structures in different data channels, for 3D images and time series.

In the Operations Window of Huygens Professional, click on Analysis > Co-localization Statistics. Refer to the program manuals for further details, or see http://support.svi.nl/wiki/ColocalizationTheory. The correspondent Scripting command is coloc.

Typically you need to execute the following steps:

1. map your 3D or 4D multichannel image to a 2D multichannel 2D image, for instance with the slice or miniMIP commands.
2. use chan2rgba to map that image to a 2D packed 4channel (RGBA) image of bytes. The chan2rgba command has a lot of options to manipulate contrast, colour and gamma. Syntax:

   source chan2rgba -> dest

3. use the -type tiffrgb option to the save command, for example:

   foo save foo.tif -type tiffrgb

See also: slice, miniMIP and chan2rgba

How to use the 'Analyze labeled image' tool that can be found in Huygens Professional?

Deconvolve first:
Thresholding noisy images is tricky since voxels from noise can be seen as objects. It's a good idea first to deconvolve your image and then find a threshold which excludes all background.

Proceed with the analysis:

1. Directly label a source image using labelDirect. Syntax:

   source labelDirect -> <destination> <threshold> <seedThreshold>
   ?-frame <frame>? ?-chan <channel>? ?-garb <volume>?

2. Do the actual analyzing: Select the ORIGINAL image and use 'Analyze labeled image' to get information about the numbered objects.
   It calculates the total volume (number of voxels in the objects, the minimum and maximum value in the objects, the sum of all voxels in the objects and the location of their Center of Mass).

   yourImage labelDirect -> b 100 150
   yourImage analyze b

The score refers to a 'probability score' that each object gets after detection. The probability is a number that is based on different types of features of objects. Objects with a high probability are most likely objects that should be tracked, while objects with a low probability are most likely 'garbage'.

The 'Optimizing object detection' screen allows you to define thresholds on the number of objects, probability, size, and intensity. You can apply these together. Objects that are above or below the thresholds will be discarded. The probability is often a good first selecting criterion to start with, since it is based on several features of structures, including brightness. If all your objects of interest are very bright and clearly distinguishable it can be that the probability selection coincide with all the bright objects.

See also Object Tracker.

This can be done in the Object Tracker after the tracks have been calculated. You then will see a window in which you can click on the 'Next' button to pop-up a track editing window. If you click in the scene on a track or on one of the 'Tracks' in the text-list, you can delete or change the tracks.

See also Object Tracker.

A complete list of the supported file formats is available here.

These functions can be found in Huygens Essential under the Tools -> Convert menu. In Huygens Professional this is listed in the Operations Window under the Convert menu. For more information on the importance of having the right image dimensions for deconvolution, please visit Convert Data Set.

Yes, there is no need to uncompress them. In Linux you can gzip all .ids files in a directory with:

gzip *.ids

or for better compression (more time-consuming):

gzip -9 *.ids

To unzip:

gunzip *.gz

The Z-direction in 3D files from Nikon confocals is not always correctly displayed unless the 'Huygens compatible' option of the EZ-C1 acquisition software by Nikon is used when saving the data. To switch on the 'Huygens Compatible' option follow these steps in the EZ-C1 software:

- Click on 'File|Save As':
- Select the '*.ids' format and press the 'Options' button.
- The second property page called 'ics' shows the 'Huygens Compatible' checkbox. Please check this option and press the 'Setup' button to fill in all the Huygens specific information.

No, they don't. Huygens writes Tiffs with the 'official' Tiff library. It is known that Tiffs written on a Mac have different byte ordering than Tiffs written on Intel. Some PC readers have trouble with that although according to the Tiff standard they should be able to handle it.

If the Tiff series is numbered with Leica style extentions the series will be read as a 4D multichannel timeseries. So instead of just a sequential numbering they should be numbered like:

psa-880_Series08_t00_z000.tif
psa-880_Series08_t00_z001.tif
...
psa-880_Series08_t01_z000.tif
psa-880_Series08_t01_z001.tif
...
psa-880_Series08_t09_z027.tif
psa-880_Series08_t09_z028.tif

Huygens Pro has a lot of conversion utilities and tools to shuffle frames and channels around. See Operations Window -> Convert

Lastly it is possible to edit the ICS file like:

layout  parameters      4
layout  order   bits    x       y       z
layout  sizes   8       256     256     64

to:

layout  parameters      5
layout  order   bits    x       y       z       t
layout  sizes   8       256     256     16      4

The only tricky thing is that the field separators must be tabs, not spaces.

Yes. After you load them, we advise you to review the microscopic parameters. You can do this in all viewers or by right-clicking on the image and selecting "Edit Parameters" or "Parameter Editor" (depending on the product).

If requested, a single channel image will be exported to a multi-tiff file. Multiple channel images will be exported to a RGB 8bit multi-tif
file (if only two channels are present the last B channel will be empty, and if more than three channels are present they will be discarded).

It also writes (and reads) as 16 bit tiff, one channel per tiff. In the case of multi-channel 3D or 4D images the Leica style suffix numbering style is used.

This happens in cases where the name ends with a number. While saving an image stack as 'tiff', Leica style numbering is used. If the name ends with a number the number will be trimmed at saving and new characters added according to the Leica style numbering: starting '000' to '019' (for a stack with 20 slices). If you wish to save your stack with numbered information add a '_' or '-' like 'decon1-'. The stack will be saved as a tiff series: decon1-000.tif, decon1-001.tif, etc.

This is not possible. The SVI ftp server is for uploading only. All available files for users are in this wiki. See e.g. the Download page.

There is a problem with compressed image data in LSM files: sometimes the compression format is not quite correct. The Huygens reader will continue reading past such defects. Since they seem to occur always at the end of the data little or no image information is lost.

Yes. Huygens is equipped with a Tiff 6.0 compatible reader which is capable of reading 16-bit tiffs.

Rename your successive planes (files) to make them a series with equal base names, like:

yourFile1.tiff -> newFile000.tif (or tiff)
yourFile2.tiff -> newFile001.tif
etc.

Huygens will read the stack as a single 3D image by selecting the image newFile.000. (Or you may skip the first N planes by opening newFileN).

In case you have multichannel images or time series you have to follow the Leica-style name convention in order to read in the tiff series as a single image. See Renaming Tiff Series and Leica Tiff.

If the particular program is able to save images in binary form or forms with a simple fixed header follow this procedure:

1. In the foreign software, save the image in binary from, for example to foo.raw. If this is not possible try saving to a fixed header format and strip the header.
2. Create an image in the Huygens System of exactly the same dimensions and microscopic parameters as the image you have saved in binary form. Save it in the default ICS format, for example foo. It creates the foo.ics/foo.ids file pair.
3. Rename foo.raw to foo.ids. This over-writes the original foo.ids but you don't need that any more.
4. Open the foo.ics/foo.ids ICS file pair in Huygens.

Save the images either as Imaris-Classic or ICS. ICS is preferred, but unfortunately Imaris does not write ICS files always correctly. The Huygens software has built-in workarounds to still read such files. It should be noted here that microscopic parameters are often missing in ICS files written by Imaris.

The current native file format for Imaris is an HDF variant. Huygens supports the HDF format as well and should not give problems when loaded in Huygens. However, Imaris stores the metadata in a separate section, which is not read properly. In some cases a black border could be added to the image, which can be cropped using the crop-tool.

The previous native file format for Imaris was 'Imaris3'. Huygens does not support the Imaris3 format directly; instead Imaris3 files, being a Tiff variant, are read in by the standard Tiff library which only recognizes the first plane.

Yes. However, Imaris Classic files do not contain all microscopic parameters needed for deconvolution. For example, the microscope type and pinhole size are missing. In case you're working with Nipkow or 2-photon systems, the excitation photon count and pinhole spacing are also absent from the file. Missing parameters are substituted by an external Tcl filter. To be on the safe side, it is always a good idea to check important parameters like sampling densities and the NA.

Yes, Huygens can export numbered TIFF series with time, z and channel index suffixes according to the Leica scheme.

Voxel values larger than 32767 are represented as negative numbers in the signed format used by the software. You can solve the problem by running some Tcl operations. Suppose you named the image with the wrap around problem `wrapped' you can correct this with the following Tcl operations:

<1> wrapped clip -> a
<2> wrapped - a -> c
<3> a convert -type float
<4> c convert -type float
<5> c * -1 -> b
<6> b max 1 -> b
<7> c + 65536 -> c
<8> c * b -> c
<9> a + c -> psf

This gets you the corrected image in float format.

The Image Cytometry Standard (ICS) was first proposed in:

• P. Dean, L. Mascio, D. Ow, D. Sudar, J. Mullikin, "Proposed standard for image cytometry data files", Cytometry, n.11, pp.561-569, 1990.

More information and source code under the LGPL license is available from: sourceforge

Read our extended explanation on how to run Huygens remotely at RemoteDisplay.

We recommend good quality hardware like HP/Compaq, IBM, Dell. We have seen problems with cheap Intel systems and not so cheap systems with AMD Athlon processors. These problems are most likely caused by inadequate power supplies and cooling.

Huygens is fully 64 bit for Windows, Mac and Linux. When possible it might be good to consider 8 GB of memory per 2 or 4 threads or cores. This relation is scalable (2-4 threads -> 8 GB, 4-8 threads ->16GB etc).

A rule of thumb for memory needs: 3 times as much FREE memory as the data-size.

Which graphical card you choose is irrelevant as all the processing by Huygens is done directly on the processors. So no need to spend money on that. Currently the Xeon processors are fast and the price performance relation is really good.

See our benchmark results.

First, it is a good idea to check how much swap space is already available with the command:

free -m

The -m switch tells free to report the space in megabytes. To increase the amount of swap space you can either allocate 1) a dedicated swap partition on disk, or 2) a swap file. Both procedures must be executed from the root account.

1) Dedicated swap partition
For this you need a dedicated free partition. This partition can be at most 2 GB in size. Suppose this partition is /dev/hdg7 then initialize the swapspace with:

mkswap /dev/hdg7

Edit the file /etc/fstab and add a new entry for this swap partition:
/dev/hdg7 swap swap defaults 0 0
Enable the new swap partition with:

swapon -a

2) Adding a new swap file
This procedure is more complex, but the advantage is that you don't need to repartition or add disks for it. First locate which of your currently mounted partitions has enough free disk space available by using the df command. Suppose you find that the filesystem which is mounted as the /mydata directory has sufficient free disk space available to hold a 2 GB swap file. Use the following commands to create it:

mkdir /mydata/swap
chmod 700 /mydata/swap

on one line:

dd if=/dev/zero of=/mydata/swap/swapfile1 bs=1024 count=2097150
chmod 600 /mydata/swap/swapfile1
mkswap -f /mydata/swap/swapfile1 2097150

The dd command may take some time to complete. After the mkswap command has completed edit the file /etc/fstab and a new single line entry for the swap file:

/mydata/swap/swapfile1  swap swap defaults  0 0

Lastly enable the new swap file with the swapon command:

swapon -a

In a default Linux installation the maximum size of sections of shared memory is too small to be of use for deconvolution. To view the current maximum size of shared memory segments run:

/sbin/sysctl kernel.shmmax

To increase this value add the following line to /etc/sysctl.conf:

kernel.shmmax = 2147483648

and then to activate this new value run this command:

/sbin/sysctl -p

To have this command run during system startup on SuSE add it to the file /etc/init.d/boot.local. RedHat systems should already look at /etc/sysctl.conf at startup. If it exists sysctl is automatically run.

First of all you need sufficient RAM and swap space. To add swap space see:

• How to add swap space on Linux?

See also:

• How do I enable the use of shared memory on Linux?

The 64 bit version of the Huygens software for Windows (XP, Vista, and 7) can address all available memory. Generic 32 bit Windows installations limit the user applications to 2 GB of memory. However, on some windows platforms there is a possibility to extend the user address space to 3GB by adding the /3GB to the boot options in the boot.ini file. The Huygens software can take full advantage of the extra 1 GB memory space.
See the following articles on Microsoft's support site:

• Main Support article on 4GB tuning: http://support.microsoft.com/default.aspx?scid=kb;en-us;291988
• Windows 2003 issues: http://support.microsoft.com/default.aspx?scid=kb;en-us;316739
• Windows NT issues: http://support.microsoft.com/kb/189293/

See also demystifying the 3GB switch.

According to the Microsoft support site, the following 32 bit platforms can support 3GB memory user space:

• Windows 2000 Advanced Server
• Windows 2000 Datacenter Server
• Windows 2003 server
• Windows XP Professional
• Windows NT Server Enterprise Edition on Intel architecture.

Basically, the way to insert the /3GB switch is to edit the files C:\boot.ini by adding one more boot line option which contains the /3GB flag.
For example, if the original boot.ini is:

[boot loader]
timeout=30
default=multi(0)disk(0)rdisk(0)partition(1)\WINDOWS
[operating systems]
multi(0)disk(0)rdisk(0)partition(1)\WINDOWS="Windows Server 2003, Enterprise" /noexecute=optout /fastdetect

you should add one more line that includes the /3GB option, i.e.

[boot loader]
timeout=30
default=multi(0)disk(0)rdisk(0)partition(1)\WINDOWS
[operating systems]
multi(0)disk(0)rdisk(0)partition(1)\WINDOWS="Windows Server 2003, Enterprise" /noexecute=optout /fastdetect
multi(0)disk(0)rdisk(0)partition(1)\WINDOWS="Windows Server 2003, Enterprise 3GB" /noexecute=optout /fastdetect /3GB

Please refer to the Microsoft support page on the boot.ini file in: http://support.microsoft.com/kb/314081

In order to view the boot.ini file on Windows XP you should open C:\ in the file browser, Choose: Tools→Folder Options. In the "Folder Options" window choose the "View" tab. Browse in the "Advanced Settings" list to "Hidden files and folders" and choose "Show hidden files and folders". Click "Apply", and you will find the boot.ini file in directory C:\.

Please make sure that there is enough virtual memory in your system. This is typically configured in: Control Panel → System → Advanced → Performance → Advanced → Virtual Memory → Change.

Open a unix shell (terminal), become root, and issue the following commands:

rm -Rf /var/lib/__db*
rpm -vv --rebuilddb

While installing a new version of Huygens Essential on my Windows machine, I got the message "Error opening file for writing: c:\Program Files\Svi\bin\essential.exe". What is wrong?

Huygens essential is already running either by your self or by someone else who was running essential before you logged him or her off in order to install the new version in your own environment.

If you tackle with any problem while using Huygens, we would appreciate it if you could send us a description of the problem, along with a debug log. This debug log is generated by Huygens, upon running in debug mode. To run Huygens in debug mode, please carry out the instructions listed here.

Yes. The embedded Batch Processor in the Huygens Software allows you to do batch deconvolution, but you will need an X-terminal or other remote graphical interface for remote usage.

HRM is a web interface to Huygens Core and allows you to control the deconvolution queue with any of the supported web browsers. Once it is properly installed in the server, no configuration is necessary for the client apart from a running browser like Firefox.

Huygens Essential and Huygens Professional have their own integrated Batch Processor, but it is more intended for single-user usage. Multiple users may run simultaneous sessions of the Batch Processor, but the multiple batch processors will compete for the same hardware resources, likely resulting in a slowdown.

HRM, on the other hand, has a queuing system intended for multiple users. Different users have their own accounts and put deconvolution jobs in the queue; they will be managed in order (first in, first out). A quota system for improved queue management can easily be implemented, and will probably be included in future versions of the HRM.

That is possible with HRM: the queue is always visible to all connected users (but users can only delete jobs they own, of course).

The Huygens Remote Manager is an open-source project and is developed by groups using Huygens at the Universities of Montpellier, Basel and Lausanne. Scientific Volume Imaging simply collaborates in its development. There is no possible fee from our side for its usage. Because it is free and open, you can adapt it to any usage you can imagine.

Still, HRM was developed with the idea of working next to the Huygens Compute Engine (HCE), to serve as a front-end to it. Moreover, the Huygens Core (an implementation of the HCE without a graphical interface and intended for large servers) is enhanced to work nicely with HRM, and it will include more and more features specially designed for a web interface like HRM.

You will indeed need a license to extend the FreeWare capabilities of Huygens Core or other SVI programs and be able to do full deconvolution, independently of whether you use it with HRM or any other means. You are always welcome to apply for a test License String!

As a summary:

• HRM is free to use and adapt.
• Huygens Core, like all the Huygens Software, has some limited FreeWare capabilities, but
• to fully do deconvolution with Huygens Core you need a License String.

As with any other package of Huygens Software, that depends on the modules and options you need. You can request a personalized quote by using this form.

The License Strings for the different Huygens Software products have a tag that specify the number of processors on which the programs are allowed to run (see License String Details). Traditionally the machines have been called "desktop" (for up to two processors) and "small", "medium", "large" or "extreme server" (for many processors).

In this sense there is no "server module". The term server refers just to a machine category for a license to run, not to extended features of the programs intended for servers.

The "server" tag in a License String refers only to the size of the machine the license is created for (see previous question). You will need it to make use of all the available processor in your system, independently of the usage of HRM.

HRM is just an interface to Huygens Core. HRM is free to use, and can save you a lot of time when programming remote-controlled batch deconvolutions. You can also adapt it freely to do not only deconvolution but any other image processing you can imagine: as an interface to the Huygens Compute Engine, many powerful Tcl Huygens commands are available.

No, HRM will work properly only with Huygens Core.

All the different products in the Huygens Suite share the same Huygens Compute Engine, therefore the deconvolution results are always the same. The difference between products is mainly in the user interface.

Both Huygens Essential and Huygens Professional have their own graphical interfaces and can not be executed without it. Therefore they can't be used with another interface like HRM on top of it.

Huygens Scripting can optionally run without a graphical interface. It requires that you program your own scripts and perhaps design your own interface. It is not intended for large scale web-based deconvolution.

Huygens Core works by default without a graphical interface and was designed to work seamlessly with HRM. Huygens Core is continuously being expanded to include more and more features specially intended for web-interfaces like HRM.

Once you have a regular web server running (including Apache, PHP and a database like PostgresQL or MySQL) installing HRM is not really difficult. You can find some guidelines based on previous experiences in Hrm Installation and you can always contact SVI for support. Remember that you will also need to install Huygens Core!

More information about the HRM installation can be found at the official website of the HRM project.

The HRM code is currently Unix-oriented, and it is primarily intended to run only in large servers running Linux. It has been tested with success in MacOS X 10.5 after a few adaptations, and it is not difficult to install because OSX comes with most of the necessary server components. It hasn't been tested on Windows but it anyone can try! Because HRM is open source code, everybody can tweak it.

Huygens Core, which was created to be used with front-end applications like HRM, is currently available for all the Huygens supported platforms (Win, Mac and Linux). If you want to give it a try, please Contact Svi.

We have installed HRM / Huygens Core without any difficulty in Fedora, Ubuntu, SuSE, Mandriva and CentOS. You really can choose what you want, all popular distributions will do as long as you install the "web server" components. (Many installations ask you about the computer main usage, and they automatically install Apache, PHP and databases if 'web server' is selected. If not, you can always install them later).

To read about which database to install see below.

HRM makes use of the database abstraction library ADOdb so many popular databases can be used in the background to store user data and image parameters. We have tested it with MySQL and PostgreSQL but in principle many others are possible, see the ADOdb project page.

Huygens Core can be installed in multiple computers and HRM can be configured to distribute tasks over these computers from a central server. (In such configuration, HRM is installed only in one server, Huygens Core in many).

HRM is prepared to handle multiple computers, to assign tasks in the queue to any free computer in the grid. In this sense, HRM provides parallel processing of images. Still, the deconvolution of each image in the queue is assigned completely to a single computer only: large images are not split among computers in the grid for parallel processing.

Because Huygens Core (as all Huygens applications) is multithreaded and uses parallel processing for its deconvolution algorithms, each image assigned to single computer will be deconvolved using parallelized procedures and will benefit from that computer having multiple processors.

You can program your own interface to Huygens Core or use other queuing application different from HRM if you want to distribute image processing tasks over the computers in the grid, with this same idea.

Since version 1.1 HRM contains previsualization tools to explore your restored results and compare them with the original images. See Hrm Screen Shots.

There's an online HRM demo installation supported by SVI, which everyone can use here.

HRM stores usage statistics of all deconvolution jobs sent to HuCore. All the statistics can be exported to a text file, which can be opened in spreadsheet processors.
To retrieve the complete HRM statistics log on to HRM as 'admin' and click on the "global statistics" icon. In the statistics panel, start and end dates can be selected to get the statistics of a particular period of time. Select a research group if you wish to get the statistics of a specific research group, otherwise select "All groups". There are several more options to select a specific statistics type, but in this case you may want to select one of the following:

• Number of jobs per user.
• Number of jobs per group.
• Microscope type.
• Total run time per user.
• Total run time per group.

Yes, the -script <myscript> option works as follows:

huscript -script myscript

All Huygens products will show a message when a new version is available. You can also check for updates manually by going to Help > About and clicking on the Check for updates button.

Deconvolution results are usually in float format, so when saved to 16-bit TIFF they sometimes need to be contrast-stretched to make the data fit in the 16-bit format. If no stretching is necessary because the data fit in the 16-bit format, there will be no scaling.

If you wish to have more control on this process you can convert the data to the desired data type before saving. This can be done with the Convert->Data and Channel type menu from the Operations Window or with the <image> convert command, e.g. a convert -type byte.

For more elaborate conversions see Manipulation->Copy&Stretch and the arithmetic menu. See also Tiff Scaling.

The syntax of the open command is:

img open <filename> ?-query? ?-foreignTo byte|int|float? ?-cmode clip|scale?

So you'd need to use the -foreignTo int -cmode clip options.

img open yourImage -foreignTo int -cmode clip

See img open.

Read our extended explanation on how to run Huygens remotely at RemoteDisplay.

Measuring distances can be done with the Twin Slicer, available in Huygens Essential and Huygens Professional in both freeware and licensed modes. Next to that the option Object Analyzer enables a full suite of vertasile analysis tools both for the whole image and parts or objects of it.

Select an image and open the Twin Slicer by either right-clicking on the image and selecting View > Twin Slicer or by the menu Visualization > Twin Slicer. In Huygens Essential there is an additional button to open the Twin Slicer in the main window.

When the Twin Slicer is opened, the image is shown in the left viewer. When you simply click on the image and then drag to another point a line a is drawn between the two points with a distance displayed next to it. In the right viewer a line plot is automatically visualized, which shows the intensity values where the line goes through. A drawn line can be deleted by selecting the line and pressing Delete on the keyboard.

You can select any view of the image by selecting a slice direction and by changing the tilt and twist, which makes it possible to measure distances is all directions.

There are quite a number of books on Tcl/Tk, below we list only the lucid book on Tcl by John Ousterhout, and the encyclopedian book written by Brent Welch et al:

• Tcl and the Tk Toolkit by John Ousterhout, ISBN: 0-201-63337-Z, Addison-Wesley, New York.
• Practical Programming in Tcl and Tk (4th Edition) by Brent Welch, Ken Jones, Jeffrey Hobbs, ISBN: 0-13-038560-3, Prentice Hall PTR.

See also Tcl Tk, Tcl Huygens, Tcl Notation.

There are many commands available, both tcl/tk commands as huygens commands. For the full list of available commands see:

• TclTkCommands

For an online tutorial and explanation about the huygens commands, see:

• Huygens Core Manual

Note: the be able to view the manual of Huygens Core you need to be logged in.