Building a light sheet microscope around an AZ100 microscope, part 2

In my previous post I talked about the basics of building a light sheet microscope from an AZ100 scope. After our initial successes with the microscope, we wanted to upgrade it to multicolor imaging and add a motorized Z stage to allow easy sample movements and potential image stitching.

For the multicolor laser source, we added a 4-line (405 / 488 / 561 / 640 nm) Vortran VersaLase laser launch. Vortran was easy to work with; since they’re located in Sacramento, they even drove up to install it for us. It’s fiber coupled; we use a doublet lens to collimate the beam from the fiber and then a cylindrical lens to produce the light sheet. A slit in front of the cylindrical lens allows trading off the width at the beam waist and the convergence of the sheet, allowing you to choose whether you want a thin sheet over a small area or a wider sheet over a larger area.

To add an emission filter wheel, we turned to Ebay. I’ve mentioned before that you can get cheap ASI parts from old GAIIx sequencers. We bought one such set of parts for the light sheet system, and then designed mounts for the filter wheel between the objective nosepiece and the filter turret, where a DIC slider housing formerly went. I’m pretty sure that this is in infinity space, and in any event the filter wheel mount isn’t that much thicker than the DIC slider housing was, so I don’t expect this to add much aberration. I had a lot of fun designing the filter wheel adapter in Autocad Inventor; I 3D printed parts of it to test the fit, and then had it machined in aluminum by proto labs. The total cost for the custom machining was around $700 and the turnaround was around a week, so I would definitely use them again.

The ASI stage mounts on the transmitted light illuminator base in place of the manual stage that had been there, via a 3D printed adapter. A second adapter mounts to the top of the plate and allows interchangeable 3D printed holders for different size cuvettes to be installed. We started with a 30 mm ID cuvette from Hellma (type 704-OG), but it was too tall to fit underneath the 5x / 0.5 objective, so we now mostly use a custom made 2 cm x 2 cm x 1 cm cuvette from FireflySci.

Files for all the custom parts are available at Github, as is the source code for the plugin we use to specify the relationship between the cuvette and objective position..

Bidirectional Z-scanning with Micro-manager and an ASI Z-stage

Conventional (unidirectional) Z-stack acquisition as compared with bidirectional Z-stack acquisition. In the conventional case, the time for the Z-stage to return to its starting position (the rescan time) limits how fast stacks can be acquired. In the bidirectional case, the stage is continuously moving, first up, then down, allowing continuous image acquisition.

Long time readers of this blog know that I’ve spent a lot of time working to make acquisition on our systems as fast as possible. Recently, I was approached with a request from Saul Kato, a new faculty member at UCSF, to go even faster. He wanted to be able to image neuronal activity in C. elegans at > 5 volumes per second.

What limits the acquisition speed of multiple volumes in Micro-manager is that it acquires Z-stacks unidirectionally and then has to return to the start position at the end of the Z-stack. This return time, also known as the retrace time, can actually add quite a bit of overhead (> 100 ms). In part, this overhead is to allow time for the piezo to return to its start position (I remember working with a Micro-manager version, many years ago, that didn’t allow enough time to return to the start, so all Z stacks after the first were missing the first plane or two). There is also some software overhead in this retrace time.

To eliminate this overhead, Saul and I set up bidirectional Z-scanning in Micro-manager. To avoid the rescan time, bidirectional Z-scanning first acquires one Z-stack ascending, followed by one descending. Because there are no large stage moves, the camera can acquire continuously during the entire process and so the overall acquisition rate is much faster.

We implemented this by taking advantage of the same trick in talking to the ASI stage that I’ve used before: Micro-manager allows you to communicate directly with the stage over the already open serial port from a script. Saul’s script then loads the ring buffer on the ASI stage with positions for both the ascending and descending Z stacks, and sets it so that camera triggers cause it to move from one plane to the next. With this set up, you just acquire a time lapse with as many frames as you want, and the Z-stacks are automatically acquired. You need to post-process the resulting stack to assemble the frames in the right order but the acquisition is very simple.

The script for doing this is on Github, as is one for turning off the bidirectional movement.

Deep-UV excitation with oblique epi-illumination

For the last several years, I’ve been working on a project to make spectrally-encoded beads using luminescent lanthanide nanophosphors [1] [2]. We use the nanophosphors to make unique spectral fingerprints for different beads by varying the concentration of different lanthanide emitters with distinct emission spectra. In particular, the nanophosphors we use are ytrrium vanadate nanocrystals doped with lanthanide emitters such as europium or dysprosium. We use lanthanide nanophosphors, rather than other fluorophores because they have narrow emission lines, are photostable, and are chemically stable. However, they have one major drawback: their excitation maximum is at 280 nm. This wavelength is so short that it is not transmitted by glass or conventional optics; instead you must use fused silica or special plastics like cyclic olefin (co)polymer to get substantial transmission. This means that conventional epi-illumination (through the objective) cannot be used to excite our samples. While there are objectives optimized for transmission of such short wavelengths, they are very expensive. Instead, for our work to date, we have used transmitted light illumination to excite our samples. However, this is relatively low brightness, illuminates a small field of view, and uses an expensive arc lamp source.

The deep-UV illuminator mounted on a 4x / 0.2 NA objective.

To try and improve on this light source, I designed an epi-illuminator for 280 nm illumination of our samples. Rather than illuminating through the objective, it consists of six deep-UV LEDs aimed at the sample. The LEDs are 280 nm Optan LEDs from Crystal IS with a ball lens to produce a narrow beam of light. Each emits ~ 3-4 mW of light, for a total of ~ 24 mW at the sample. They are mounted in a 3D-printed mount designed to aim each LED at the focal point of the lens. Clean up filters are mounted in front of each LED. These are 300 / 80 nm bandpass filters from Semrock, custom cut to 9mm diameter. Continue reading


  1. R.E. Gerver, R. Gómez-Sjöberg, B.C. Baxter, K.S. Thorn, P.M. Fordyce, C.A. Diaz-Botia, B.A. Helms, and J.L. DeRisi, "Programmable microfluidic synthesis of spectrally encoded microspheres", Lab Chip, vol. 12, pp. 4716-4723, 2012.
  2. H.Q. Nguyen, B.C. Baxter, K. Brower, C.A. Diaz-Botia, J.L. DeRisi, P.M. Fordyce, and K.S. Thorn, "Programmable Microfluidic Synthesis of Over One Thousand Uniquely Identifiable Spectral Codes", Advanced Optical Materials, vol. 5, pp. 1600548, 2016.

Building a Microscope from Thorlabs Parts

I meant to document this better, but now it doesn’t look like I’ll have time to do so. This is a scope I assembled from Thorlabs parts for a research project involving tracking bead movement by laser deflection. We ended up using a different assembly eventually, but I thought I would put up the parts list here in case it’s useful to anyone.

thorlabs scope


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New Sutter LED combiner

Sutter recently announced a clever new LED combiner that allows you to combine multiple LED light sources into a single output beam. It relies on the fact that interference filters are very good reflectors of the wavelengths they don’t transmit, so you can use a filter to simultaneously pass the output of one LED while reflecting wavelengths from other LEDs. The concept is shown in the diagram below:

The Sutter 421 LED combiner. The number at each position indicates how many reflections that LED undergoes before reaching the output. Image courtesy of Sutter.

One of the many nice things about this concept is that changing LED wavelengths is really easy: you just replace the LED and the filter in front of it. You can also mount a second pentagon on the first to combine up to seven wavelengths (in principle you could even cascade a third pentagon to get 10 wavelengths, but at some point the filter designs get pretty tricky and the losses add up). You can also combine light sources other than LEDs, provided you can find appropriate interference filters.

We demoed a six color version of this a few weeks ago, using two pentagons and LEDs for DAPI, FITC, Cy3, Cy5, CFP, and YFP. We tested it with a Semrock Sedat Quad filter set and Chroma GFP/RFP and CFP/YFP filter sets. At all wavelengths tested it was as bright or brighter (in some cases as much as 10-fold brighter) than the Lambda XL we were using as a reference.  We’re now working with Sutter to get a seven color version of this (including 340 nm excitation for Fura-2) to install on our microscope. This will allow us to synchronize the LEDs to the camera, so that the LEDs are only on when the camera is exposing, minimizing photobleaching and phototoxicity. This should be a very nice LED illumination option for microscopy, particularly for users who want a modular system that’s easy to modify as needed.

Testing a Point Grey Camera for Fluorescence Microscopy

About two years ago, I mentioned Point Grey cameras. These are cameras sold to the machine vision and industrial inspection market, and are much cheaper than typical microscopy cameras – most are <$1000. Point Grey puts out very nice spec sheets listing all of their cameras, and the specifications for some are pretty impressive – cameras with < 3e- read noise for ~$500. Nico Stuurman has recently written a Micro-manager driver for these cameras, and was kind enough to let me test one of these cameras. We mounted it opposite a Hamamatsu Flash4.0 (an older Flash4.0, with ~72% QE), and did a qualitative comparison by taking sequential images of the same test slide on both cameras.

The Point Grey camera we tested was a Chameleon3 CM3-U3-31S4M. This uses a Sony IMX265 sensor, which has 2048 x 1536 3.45 μm pixels, with 71% QE, <3e- read noise, and sells for ~$500. It can run at up to 55 fps. On paper, this camera should perform almost as well as the Flash 4.0. The images below are of a Texas red-phalloidin stained cell, captured with a 20x / 0.75 NA objective and a 10 ms exposure on both cameras. Click on the images to see the full size image.


The Flash 4.0 camera, 10 ms exposure. Click for full size.


The Point Grey camera, 10 ms exposure. Click for full size.

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New Nikon Stand

Nikon has just announced a new stand, the Ti2.  Some noteworthy features, including a 25mm camera port with an F-mount (with a new tube lens and larger filter cubes; it looks like the Plan Apo λ objectives are flat across this field), an LED brightfield illuminator with a fly-eye lens for uniform illumination, a motorized correction collar, an internal camera for back focal plane imaging, and encoding of all microscope components.

Building a light sheet microscope around a Nikon AZ100, Part 1

A few years ago we got a Nikon AZ100 microscope on indefinite loan from a lab here that no longer was using. The AZ100 is an interesting microscope – it has low magnification objectives with relatively high numerical apertures (we have 1x / 0.1, 2x / 0.2, and 5x / 0.5 objectives) combined with a 1x – 8x optical zoom system to allow both large field-of-view imaging and high resolution imaging of the same sample. I initially set this up for routine fluorescence imaging, but it didn’t fill a useful niche and so largely went unused.

As groups on campus began testing various tissue clearing methods (CLARITY [1], PACT [2], iDISCO [3], …), I realized that this would make a good base for a simple “Ultramicroscope”-style [4] light sheet microscope. This is about the simplest kind of light sheet microscope you can build; you simply use a cylindrical lens to reshape an expanded laser beam to a sheet that propagates perpendicular to the optical axis of the microscope.  We had an old 561 nm Coherent Sapphire laser sitting around from a rebuild of the laser launch on our spinning disk confocal, so a few hundred dollars in Thorlabs parts sufficed to set up a demo system. The sample is placed in a cuvette on the microscope stage, illuminated with the light sheet from the side, and imaged with the objective from above.

The initial light sheet test system.

The initial light sheet test system. The laser is mounted on the black table; to the left you can see the mirrors used to direct the beam to propagate through the image plane, perpendicular to the optical axis. The cage system holds a Galilean beam expander and a slit; the cylindrical lens sits inside the dark enclosure. In the inset you can see the cylindrical lens and fluorescence excited in an agarose cylinder doped with fluorescent beads.

Continue reading


  1. K. Chung, J. Wallace, S. Kim, S. Kalyanasundaram, A.S. Andalman, T.J. Davidson, J.J. Mirzabekov, K.A. Zalocusky, J. Mattis, A.K. Denisin, S. Pak, H. Bernstein, C. Ramakrishnan, L. Grosenick, V. Gradinaru, and K. Deisseroth, "Structural and molecular interrogation of intact biological systems", Nature, vol. 497, pp. 332-337, 2013.
  2. B. Yang, J. Treweek, R. Kulkarni, B. Deverman, C. Chen, E. Lubeck, S. Shah, L. Cai, and V. Gradinaru, "Single-Cell Phenotyping within Transparent Intact Tissue through Whole-Body Clearing", Cell, vol. 158, pp. 945-958, 2014.
  3. N. Renier, Z. Wu, D. Simon, J. Yang, P. Ariel, and M. Tessier-Lavigne, "iDISCO: A Simple, Rapid Method to Immunolabel Large Tissue Samples for Volume Imaging", Cell, vol. 159, pp. 896-910, 2014.
  4. H. Dodt, U. Leischner, A. Schierloh, N. Jährling, C.P. Mauch, K. Deininger, J.M. Deussing, M. Eder, W. Zieglgänsberger, and K. Becker, "Ultramicroscopy: three-dimensional visualization of neuronal networks in the whole mouse brain", Nature Methods, vol. 4, pp. 331-336, 2007.

High Speed PCIe SSDs

Those of you who’ve been reading this blog since it’s inception will remember that I used to post a lot about solid state drives, because we spent a lot of time trying to handle the 1 GB/sec bandwidth of sCMOS cameras back in 2013. We standardized on RAID 0 arrays of four Samsung Pro SSDs, and I stopped thinking about it, because that was good enough for our purposes.

Since then, however, quite a bit has changed. You can now get an 512 GB SSD card that can write at 1.5 GB/sec and read at 3 GB/sec (the Samsung Pro 950) for $350. Newer Samsung products promise slightly faster read / write speeds at disk sizes up to 1 TB. These use the M.2 interface, designed for SSDs, but PCIe to M.2 adapters are readily available if your motherboard doesn’t have an M.2 slot. To get full speeds you’ll need a motherboard with a PCIe 3.0  x4 slot available.

Another thing that’s changed is that sCMOS cameras have made it really easy to capture large data sets. In the last year people really seen to have taken advantage of this and we’re seeing a lot of people acquiring 100GB+ data sets, and often up to 1 TB. A big consequence of this is that data processing is increasingly becoming I/O bound. A mechanical hard drive tops out at around 120 MB/sec sequential read/write speed. At those speeds, just reading a 20 GB file takes around 3 min. We’ve seen this become a major issue where exporting a 500 GB data set from Nikon ND2 to TIFF takes hours.

If you’re doing any processing of large data sets, it’s very advantageous to have a fast drive like this to speed up I/O. There’s still a challenge in getting data on and off of these drives, since gigabit ethernet tops out at around 100 MB/sec, and most USB3 drives max out around 250 MB/sec (as an aside, the Samsung T3 looks pretty promising, with 450 MB/sec transfer rates). Finally, most people (at least here) still want their data on a mechanical drive for long term storage. But at least once you have data on the fast drive the I/O bottleneck gets better by about 10 – 20-fold.

All of this would be well worth thinking about if you’re building the compute environment for a microscopy core from scratch. I imagine you’d want fast local storage for data processing, 10 Gbit networking to move data around, and some kind of slow archiving to long term storage. You’d also want to educate people about data handling so that they think about these issues when designing their experiments.