Category Archives: Uncategorized

IN FOCUS: The ‘3D Pollen Library’: An Update

Above: Transparent 3D rendering of a dandelion (Taraxacum officinale) pollen grain. Surface exine displayed in green, inner intine structure in blue. Image produced by Dr Anthony Hayes, Bioimaging Hub, Cardiff School of Biosciences.

Over the last 12 months I’ve been working closely with biovisualisation specialist Dr Kristen Brown at NIH3D to curate our 3D pollen model resources into a purpose-built 3D collection:  the ‘3D Pollen Library’. This collection is now featured on the NIH3D homepage and, at the time of writing, represents the largest collection of 3D pollen grain models worldwide. To date, it contains over a hundred entries together with taxonomic metadata and links to other well-established online pollen resources – a significant achievement considering its humble beginnings (you can read more about the background work leading up to it here.)

Enormous thanks must go to Kristen and her team who have been incredibly helpful throughout in accommodating my (many) requests regarding the ‘look and feel’ of the curated collection. They have done an absolutely splendid job.

Above: A small selection of surface rendered pollen grains in the 3D Pollen Library collection. These examples were recently created in collaboration with Dr Heather Pardoe, Amgueddfa Cymru, National Museum Wales.

Above: the user interface for visualisation and manipulation of the 3D pollen models. The models are all viewable  as surface rendered or wireframe meshes and can be downloaded in x3d, stl, glb and wrl file formats for 3D printing and AR/VR visualisation. The source confocal data is also available for download, as is the published methodology for creating the models.

In parallel with the above, I’ve established an ongoing collaboration with Dr Heather Pardoe (senior botanist and chief palynologist at Amgueddfa Cymru, National Museum Wales). This collaboration has allowed us to digitise and 3D model pollen grains and spores from selected plant species in the museum’s extensive archival pollen collection using methodology we’ve developed in-house at the Bioimaging Hub. The 3D pollen models produced via this collaboration will be added to our NIH3D library as a separate pollen sub-collection, as well as being viewable as part of the full collection. It is envisaged that these models will have significant utility as educational tools for teaching and exhibition.

Further reading:


NEWS: Updated Covid Rules: Resumption of Hands-on Support and Training.

A huge and heartfelt thank you to all users and support staff of the Bioimaging Hub for your strict adherence to our covid security measures over the last 12 months. It has been an extremely difficult year for all of us and we have tried to manage the situation as effectively and as safely as possible, working within the security framework provided by Cardiff University and Welsh government.

In line with the latest guidance, we are pleased to now begin relaxing some of our covid security measures and to be in a position where we can reintroduce direct hands-on support and training for our microscope systems.  To facilitate this provision, it remains vitally important that users follow the new guidance protocols, as detailed below.

Before entering the Bioimaging Hub:

  • All users must familiarize themselves with current Welsh Government Coronavirus (Covid-19) Guidance and read the Bioimaging Hub’s updated Coronavirus risk assessment.
  • Users must not visit the Bioimaging Hub if they are displaying any symptoms of Covid-19; if they have been in a high risk area; or had recent contact with a covid-positive individual without confirmation of a negative test result.
  • All users are advised to make regular use of the Cardiff University covid screening service  or to utilise rapid flow testing methodology that is now widely available through the NHS and pharmacies.

Room occupancy status and technical support/training:

  • Room occupancy status has now increased to two independent users/two research bubbles per microscopy suite, up to a maximum of four people per microscopy suite in total.
  • Direct hands-on training and support for our microscope systems will now resume under social distancing rules and with PPE including face coverings.

COVID working Regulations within the Bioimaging Research Hub:

  • Users should continue to use the room booking calendars and should specify which microscope system they require in the title section of the booking request (i.e., specify user name and microscope system). Booking details here.
  • Social distancing rules remain in place within all areas of the Bioimaging Hub.
  • PPE remains mandatory including the use of face coverings in all communal areas.
  • Microscope cleaning procedures before and after use remain mandatory.
  • Histology sample drop-offs and collections should continue to be arranged via email (


AJH 28.9.2021

IN-FOCUS: Brushing Up On Your Background Knowledge.

With the dizzying pace of technological innovation and ongoing advances in microscopy and imaging, it is becoming increasingly difficult to keep abreast current developments in the field. With this in mind, we’ve rounded up some essential resources, from basic to advanced, that will keep you informed and updated.

Online magazines & journals:

Educational portals:


NEWS: Reopening of the Bioimaging Research Hub: New COVID 19-Security Measures.

For the time being, use of the Bioimaging Hub is going to be a very different experience for everyone. In drawing up these guidelines we’ve taken a lead from the Welsh Government and have started with a very cautious approach that we’ll constantly review and relax where possible. For now, the guidance may seem quite stringent so please bear with us while we all adapt to this new way of working.

Access is not permitted to the Bioimaging Research Hub without (i) completing a risk assessment form that should cover all planned activities within the facility and (ii) reading all relevant supporting information (see below). A template risk assessment is available through the Bioimaging Hub’s main web pages.

Before completing the risk assessment form, all users must download and read the following documents:

Cardiff University’s COVID-19 secure organisational risk assessment

The Bioimaging Hub’s local COVID-19 security guidelines

DO NOT enter the Bioimaging Research Hub:

  • If you are SARS-Cov-2 positive or are displaying symptoms of COVID-19 infection (e.g. persistent coughing, elevated temperature, anosmia, sickness).
  • If you have been in a high-risk area or have had recent contact with confirmed SARS-CoV-2 positive individuals within the last 14 days.
  • If you have an underlying health condition and are concerned it will put you at greater risk of developing severe COVID-19 symptoms. N.B. essential work can be undertaken by Bioimaging Hub staff at a supported rate if necessary.

COVID Working Regulations within the Bioimaging Research Hub

All users of the facility:

  • MUST read the above documentation before entering the Bioimaging Research Hub.
  • MUST contact before entering the facility.You are not permitted to simply drop-in unplanned. Drop off/collection of histology samples must be arranged in advance. Researchers must knock before entering histology via E/0.08 and observe all safety directives outlined in this document. Users must complete a histology request form to specify processing preferences in advance of their visit.
  • MUST wash their hands upon entering and leaving the facility (hand wash station opposite office area). Multiple gel hand wash dispensers are also available within the microscopy suites.
  • MUST wear appropriate personal protective equipment (PPE) within the facility, including a buttoned-up lab coat and gloves (gloves are provided in all microscopy suites). Eye protection is advisable.
  • MUST clean the imaging equipment and immediate working area before and after use (cleaning instructions are available at each microscope station; alcohol spray and lab roll have been provided for this purpose).
  • MUST observe social distancing (i.e. 2 metres inter-personal space) within the Hub. The main corridor should be used by only one person at a time. While social distancing measures are in place no training is available and technical support will be provided on a remote basis via telephone (main office: 02920876611; shared office: 02922510220; histology: 02920875139).
  • MUST NOT enter the staff office area (0.14A) or histology suites (E/0.06-E/0.07). Drop off/collection of histology samples must be arranged in advance (use E/0.08).
  • MUST knock before entering any room. All microscopy suites are now single occupancy (i.e. one in, one out with at least 15 mins user-free time between bookings). The microscope booking calendars have been replaced with room booking calendars, as follows: “BIOSI – E/0.03 – Confocal/Lightsheet microscopy”, “BIOSI – E/0.04 – widefield microscopy”, “BIOSI – E/0.05 – spinning disc microscopy”. New booking instructions can be found here.
  • MUST set the room occupancy status (vacant/in use) on the door sliders of the microscopy suites before and after use.


German BioImaging recommendations for operating Imaging Core Facilities in a research environment during the SARS-CoV-2 pandemic

Leica: How to sanitize a microscope

Olympus: How to clean and sterilize your microscope

Zeiss: Cleaning and disinfecting the microscope and its optical components

IN FOCUS: Bioimaging Hub’s YouTube Channel – Rebooted.

Above: the new look Bioimaging hub YouTube channel.

With the University (and entire planet) in lockdown due to the ongoing covid 19 pandemic, now seems as good time as any to make you aware of the Bioimaging hub’s ‘rebooted’ YouTube channel, if you was not already aware of it.

A decade is a long time in imaging. Way back in 2009 we set up a YouTube channel to showcase the capability of our new, all-singing, all-dancing Leica SP2 confocal microscope. At the time, we uploaded a collection of short 3D animation sequences that highlighted some of our ongoing research applications. Fast forward eleven years. Whilst many of those early demo videos have been highly viewed (one over 20 thousand times) they’re starting to look rather dated, particularly when compared to the material that we’re now producing using our new confocal and lightsheet systems and 3D analysis software. As I say, a decade is a long time in imaging.

So, with the advent of spring, we thought it was high time we dusted down our YouTube channel and gave it an overhaul. As well as introducing lots of nice new image content from some of our latest 3D imaging systems, we felt that the channel would have a greater sense of purpose if we were to develop it into an educational resource for microscopy and bioimaging, with obvious relevance for remote learning (i.e., perfect in our current circumstances). Consequently, we have started to collate the most useful and relevant of YouTube’s microscopy-related content (webinars, tutorials, demonstrations etc), ranging from the basic principles of light microscopy to cutting-edge fluorescence-based nanoscopy techniques such as FLIM, so that they are all placed under one roof for your convenience : )

One of the things we hoped to provide our userbase was a series of video tutorials for the hub’s many microscope systems. Whilst there’s a great deal of useful training material on YouTube, in the main, it tends to be aimed at many of the high-end, turn-key imaging systems. Furthermore, not all microscopes are created equal, they each have their own peculiarities which reflect their intended function and most ‘evolve’ over time, through upgrades, to accomodate the vagaries of research. So, with this in mind, we have started to create our very own bespoke training videos for each of the hub’s microscope systems (example here).

The new training videos will supplement the standard operating procedures (SOPs) we have written for all of our imaging equipment and should provide an invaluable resource for user training and e-learning. As such, they will be embedded within the appropriate sections of the hub’s SOP repository (read more here). The online video content and their associated SOPs will be viewable at the click of a mouse button via desktop shortcuts on all of our microscope-associated PCs allowing easy access during instrument operation.

If you have time on your hands, then please pop over to YouTube and take a look at how our channel is developing (link here). It’s still work in progress but, as I say, it has the potential to be an extremely useful resource; not only for hub users, but anyone with a passing interest in microscopy and bioimaging. Constructive feedback is welcomed.


CORE EQUIPMENT: Widefield Microscope Upgrades.

The Bioimaging Hub’s conventional widefield microscope systems have recently received some performance upgrades. Details of upgrades below:

N.B. All of the above systems are now networked via 1GB desktop switches. Up-to-date standard operating procedures and risk assessments for each system are available through the Bioimaging hub’s online SOP repository via their desktop folders (‘Read me before use’). See a member of staff for further details.


IN FOCUS: Winging It In Paleobiology: Strange Tails from a Strange Time.

Above: Some of the exhibits on display, including the Meganeura model made by the Bioimaging Hub, at ‘The Fossil Swamp’ exhibition at Cardiff Museum.

How do you make a dragon-fly (ask it nicely, I suppose)? Well, this was the question we were asking ourselves a few weeks ago after an email enquiry from Dr Trevor Bailey of the National Museum of Wales. Trevor is one of the museum’s senior paleontologists and is involved in curating many of the museum’s public exhibitions and programmes involving fossils and prehistoric life (You can read more via his profile page here).

Trevor had contacted us to enquire whether we might be able to help with a forthcoming exhibition at the museum, called ‘The Fossil Swamp‘, by making a scale replica model of an extinct insect species similar in appearance to a modern dragon fly (it’s actually classed as a Griffin fly), but with one big difference (and I mean BIG): its size. How big I hear you say? Well, to give you an idea of its sizeable dimensions, the wingspan of Meganeura was approximately 0.7 metres long (i.e. roughly the same wing span of a large, adult sparrow hawk)! Indeed, this is how the insect came to earn its rather ominous sounding moniker: ‘mega-neura’ means ‘large-nerved’, referring to the network of large veins supporting the insects enormous wings (Brongniart, 1893)

In fact, Meganeura monyii is one of the largest known flying insect species ever to grace planet earth. It lived more than 300 million years ago, in the carboniferous period where the atmospheric oxygen concentration of air was much higher than that of today (around 35% then, instead of 21% now) which, it is thought, allowed the insects of that period to grow to enormous proportions (insects breathe through small holes, or ‘spiracles’ in their body walls connected to branched air tubes called ‘tracheoles’ which convey oxygen to their internal tissues). Furthermore, at the time Meganeura was buzzing about, bugging the primitive lifeforms of the day, there were no other aerial vertebrate predators around (in fact, birds arrived to the table 75 million years later) so it could pretty much act with total impunity!

So Trevor supplied us with a digital model of the insect for 3D printing, together with the desired dimensions, based on recorded fossil evidence (Brongniart, 1893). Interestingly, the digital mesh was actually created as a component of a carboniferous forest simulation  by a colleague of his in Germany  (link here).  In order for us to 3D print Meganeura’s body to scale using our Ultimaker 3 extended 3D printer, we had to fabricate the head and thorax separately to the abdomen and then re-attach these after removal of their supporting scaffolds. We printed these using polylactic acid (PLA) filament at an intermediate print resolution. The first attempt looked okay, but the finished model was rather blocky in appearance, so we smoothed the digital mesh and then reprinted at a higher resolution with much better results.

1-2 digital reconstructions of Meganeura monyii; 3-5 3D prints of the separate body sections; 6 Support scaffolds removed and body sections reunited – note block appearance of model. The digital mesh was filtered and the model reprinted with much better results (see below).

The next challenge was those huge wings. I downloaded .png image files of the venation patterns recorded by Brogniart (1893) here. However they were simply too large to 3D print at the desired thickness (and believe us, we tried) so a different approach was necessary. Each wing was laser printed onto a separate sheet of acetate. These were then cut out and laminated – the composite structure increasing the rigidity of the wing but still allowing realistic flexion. To attach the wings to the thorax, I drilled holes through adjacent thoracic segments and fed lengths of wire through the holes to support the leading edge of each wing pair. The wire was then bonded in place to prevent any lateral displacement.

7 Wing venation pattern after Brogniart (1893) downloaded from the web; 8: wings laser printed to scale on acetate and laminated; 9 laminated wings cut to final shape.

Next up was the paint job, which became a labour of love (and exercise in mindfulness) in my spare time! Now, unfortunately, no one knows what colours or patterns adorned the body surface of Meganeura as the fossil evidence is all black and white. Artist’s impressions are therefore based loosely on modern equivalents (e.g dragonflies, damsel flies etc), or have just been made up to make the insect look as fearsome as possible – it was a carnivorous predator after all! So after a few Google image searches, just to get some ideas, I finally went with a black and yellow/orange colour scheme with iridescent bronze eyes (with artistic input from Trevor and my daughters). I used acrylic paints, purchased cheaply from The Works, which gave good adhesion and cover without the necessity of a primer coat. Fine detail was added under magnified optics.

10 Smoothed model with base coat of acrylic. Wires have been inserted through thoracic segments to support leading edge of wing pairs; 11-13 Model has been painted and the first pair of wings are awaiting attachment.

When the model was fully painted we attached the wings to the wire frames using extra strong clear sellotape before taking it over to Alexandra Gardens, opposite the School of Biosciences, for some wildlife photography, doing our best not to frighten the native fauna (or general public)!

14-15 The finished Meganeura model – a ferocious looking beast!
16-17 …and doing its best to blend in with the local flora!

The model will be on display as part of the Fossil Swamp exhibition in the National Museum of Wales at Cardiff from 18th May, 2019 to 17th May, 2020 along with lots of other amazing artefacts from the carboniferous period. Please go along to visit – it promises to be a fantastic family day out.


Further reading

  • Brongniart (1893) Recherches pour servir á l’histoire des insectes fossiles des temps primaires : procédées d’une étude sur la nervation des ailes des insectes (Research to serve the history of fossil insects of the early ages : preceded by a study on the wing venation of insects).


Digital mesh of Meganeura monyii was taken from the Carboniferous forest simulation, page author and domain holder Heiko Achilles; 3D printing by Dr Pete Watson; Model painting, wing fabrication and finishing by Dr Tony Hayes; Wildlife photography by Marc Isaacs. Blog post by Dr Tony Hayes.

IN FOCUS: Immersive Microscopy – 3D Visualisation and Manipulation of Microscopic Samples Through Virtual Reality.

Above: The view inside our Oculus Go VR headset: getting some top-spin on some of our 3D pollen grains!

Hands up who’s seen the provocative Stephen Spielberg sci-fi thriller Minority Report? In the movie, the main protagonist, chief of ‘pre-crime’ John Anderton played by Tom Cruise, investigates a future crime via a cool gesture-based holographic virtual reality (VR) interface. Whilst current VR technology isn’t quite that far into the future, it’s certainly not far off. Indeed, virtual reality is now becoming a reality in microscopy as researchers strive to improve their 3D understanding of complex biological samples. As creator of both the confocal microscope and the head-mounted display, a forerunner of the VR headset, Marvin Minsky would certainly approve of the convergence of these two technologies. The potential is enormous: imagine, for example, being able to take a virtual tour inside a tumour, to climb into an intestinal crypt or to peel apart the posterior parietal cortex – and all without getting your hands dirty!

‘Immersive microscopy’, as it is now known, is an area of imaging in which Zeiss in partnership with software developers arivis are currently leading the field (you can learn more here). To get in on the act, the Bioimaging Research Hub at Cardiff School of Biosciences has been developing a VR application of our own for visualisation and manipulation of volume datasets generated by the Hub’s various 3D imaging modalities. We anticipate that this technology will have significant relevance not only to imaging research within the school, but also to teaching and science outreach and engagement.

We’ve been using the affordable Oculus Go VR standalone headset and controller in association with the Unreal 4 games engine to create VR environments allowing interaction with our whole range of surface rendered 3D models. These range from microscopic biological samples imaged by confocal or lightsheet microscopy, such as cells or pollen grains, to large, photo-realistic anatomical models generated via photogrammetry.

As proof of principle we’ve developed a working prototype that allows users to manipulate 3D models of pollen grains in virtual space. You can see this in action in the movie above. We’re planning further developments of the system including new virtual 3D environments, different 3D models and object physics, and features such as interactive sample annotation via pop up GUIs. The great thing about VR of course is that we’re limited only by our imagination. To borrow a quote from John Lennnon, if ‘reality leaves a lot to the imagination’ then VR leaves a lot more!


Further reading


IN FOCUS: Standard Operation Procedures (SOP) Repository.

Above: A screenshot of the Bioimaging Hub’s SOP repository

If you wasn’t already aware of the Bioimaging Hub’s SOP repository (N.B. there are shortcuts set up on all of the networked PCs within the facility), then please take a look at your earliest convenience. The database was set up as a wiki to provide Hub users with up to date protocols and tutorials for all of our imaging systems, experimental guidelines for sample preparation, health and safety information in a variety of multimedia formats in one convenient and easily accessible location. It’s still  work in progress and we would welcome any feedback on how the resource could be further developed or improved.

AJH 7.1.19

IN FOCUS: Imaging Cleared Tissues by Lightsheet Microscopy.

We’ve had a Zeiss Lightsheet Z.1 system in the Bioimaging Hub for a little while. In the main, the system  has been used to examine small developmental organisms (e.g. zebrafish larvae) and organoids that can be introduced into the lightsheet sample chamber via thin  glass capillary tubes (0.68-2.15mm diameter) or via a 1 ml plastic syringe.  This is accomplished by embedding the sample in molten low melting point agarose, drawing it into the capillary tube/syringe and then, once the agarose has set,  positioning the sample into the light path by displacing the solid agarose cylinder out of the capillary/syringe via a plunger.

To support a new programme of research, the Bioimaging Research Hub recently purchased a state of the art  X-CLARITY tissue clearing system. This allows much larger tissue and organ samples to be rendered transparent quickly, efficiently and reproducibly for both confocal and lightsheet microscopy. Unfortunately, due to their larger size, the samples cannot be introduced into the lightsheet sample chamber via the procedure described above.  In this technical feature we have evaluated a range of procedures for lightsheet presentation of large cleared mammalian tissues and organs.

The test sample we received in PBS had been processed  by a colleague using the X-CLARITY system using standard methodologies recommended by the manufacturer. The tissue was completely transparent  after clearing, however its transferal to PBS (e.g. for post-clearing immuno-labelling) resulted in a marked change in opacity, the tissue becoming cloudy white in appearance. We thus returned the sample to distilled water (overnight at 4oC) and observed a return to optical clarity with slight osmotic swelling of the tissue.

Above: The cleared sample has a translucent, jelly-like appearance.

Generally, cleared tissue has a higher refractive index than water (n=1.33) and  X-CLARITY tissue clearing results in a refractive index close to 1.45. To avoid introducing optical aberrations that can limit resolution,  RI-matching of substrates and optics is recommended.  Consequently, our plan was to transfer the cleared tissue into X-CLARITY RI-matched (n=1.45) mounting medium and set up the lightsheet microscope for imaging of cleared tissues using a low power x5 detection objective (and x5 left and right illumination objectives) which would allow us to capture a large image field. 

Prior to fitting the x5 detection objective an RI-matched spacer ring (see below) was first screwed into the detection objective mount.

Above: Spacer ring for n=1.45 lenses.

After the n=1.45 spacer ring was fitted, the x5 detection objective was screwed into place (seen centrally in below image) followed by the x5 illumination objectives to the left and right (see below).

Above: Light sheet objectives. Illumination on left and right, observation to the rear.

Once the objective lenses had been screwed into place, the sample chamber was inserted (see below). The use of a clearing mountant requires a specific n=1.45 sample chamber. We used the n=1.45 chamber for the x5 (air) detection objective. This chamber has glass portals (coverslips) on each of its  vertical facets (unlike the x20 clearing chamber that is open at the rear to accomodate the x20 detection lens designed for immersion observation).

Above: Sample chamber for clearing (n=1.45).

Unfortunately, as it turned out, the RI-matched X-CLARITY mounting medium for optimum imaging of X-CLARITY cleared samples wasn’t available to us on the day, necessitating a quick re-think. As the tissue sample remained in distilled water  we decided to image, sub-optimally, in this medium. To do this we quickly swapped the n=1.45 spacer ring on the detection objective to the n=1.33 spacer. We then switched to the standard (water-based) sample chamber.

With the system set-up for imaging we set about preparing the tissue sample for presentation to the lightsheet.  As mentioned earlier, large tissue samples cannot be delivered to the sample chamber from above, as the delivery port of the specimen stage has a maximum aperture of 1cm across. This necessitates (i) removing the sample chamber (ii) introducing the specimen holder into the delivery port, (iii) manually lowering the specimen stage into place (iv) attaching the sample to the specimen holder (v) manually raising the specimen stage, (vi) re-introducing the sample chamber, and then (vii) carefully lowering the specimen into the sample chamber which can then (viii) be flooded with mounting medium.

The initial idea was to present the sample to the lightsheet, as described above, by attaching it to a 1ml plastic syringe. The syringe is introduced into the delivery port of the lightsheet via a metal sample holder disc (shown below).

Above: Holder for 1mm syringe.

The syringe is centred in the sample holder disc via a metal adaptor collar (shown below), which must be slid along the syringe barrel all the way to its flange (finger grips). When we tried this using the BD Plastipak syringes supplied by Zeiss we found that the barrels were too thick at the base so that the the collar would not sit flush with the flange!

Above: Metal collar for the syringe holder.

In order to make it fit, we carefully shaved off the excess  plastic  at the base using a razor blade.

Above: Carefully shaving plastic off the syringe to make it fit.

This allowed the adaptor collar to be pushed flush against the barrel flange (see below).

Above: Syringe with the adaptor collar in the correct position.

The flanges themselves also required a trim as they were too long to position underneath the supporting plates of the specimen holder disc. Note to self: we must find another plastic syringe supplier!

Once the syringe had been modified to correctly fit the sample holder it was introduced into the delivery port of the specimen stage, in loading position, by aligning the white markers  (see below)   

Above: Syringe plus holder inserted into the delivery port of the sample stage (note correct alignment of white markers).

With the front entrance of the lightsheet open and the sample chamber removed, the stage could be safely lowered via the manual stage controller with the safety interlock button depressed (see below). 

Above: Button for safety interlock under the chamber door.

The stage was lowered so that the syringe tip was accessible from the front entrance of the lightsheet (see below).

Above: Syringe dropped down to an accessible position.

Our first thought was to impale the tissue sample onto a syringe needle so that it could then be attached to the tip of the syringe (see below).

Above: Using a needle to impale the sample.

However, this approach failed miserably as the sample slid off the needle under its own weight.  In an attempt to resolve this problem, a hook was fashioned from the needle in the hope that this would support the weight of the tissue (see below).

Above: A pair of pliers was used to bend the needle and make a hook.

Unfortunately, this approach also failed, as the hook tore through the soft tissue like a hot knife through butter.

We decided therefore to chemically bond the tissue to one of the short adaptor stubs included with the lightsheet system with super-glue. The adaptor stubs can be used with the standard sample holder stem designed for capillary insertion. They attach to the base of the stem via an internal locking rod with screw mechanism (shown below).

Above: Sample stub for glued samples.

To introduce the sample holder stem into the sample chamber (see below) we used essentially the same process as that described above for the syringe.

Above: The sample holder stem being  lowered into position (the central locking rod can be seen protruding out)

The tissue sample was carefully super-glued on to the adaptor stub for mounting onto the sample holder stem.

Above: Super-gluing the sample on to the adaptor stub.

Again, under its own weight, the sample tore off the stub leaving some adherent surface tissue behind (see below)

Above: Adaptor stub with tissue torn off.

It seemed pertinent at this stage to reduce the sample volume as  it was clearly the weight of the tissue that was causing it to detach. Having already visualised the sample under epifluorescence using a stereo zoom microscope we had a very good idea of where the fluorescent signal was localised in the tissue. Consequently, we  reduced the sample to approximately one third of its original size and again glued it to the sample stub ensuring that the region of interest would be accessible to the lightsheet (see below).

Above: Sample cut down to size and attached successfully to stub.

This time it held. The adaptor stub was then carefully secured to the sample holder stem via its locking rod, the stage manually raised  and the sample chamber introduced into the lightsheet. The sample was manually lowered  into position so that it was visible through the front viewing portal of the sample chamber. The sample chamber was then carefully filled with distilled water for imaging (remember, we didn’t have any X-CLARITY mounting medium at this stage).

Above: Sample positioned in the imaging chamber.

With the sample in place, we then set up the lightsheet for imaging GFP fluorescence. Due to the large sample size, we found that we could only image from one side (the lightsheet couldn’t penetrate the entire sample without being scattered or attenuated). The sample was therefore re-oriented so that the region of interest was presented directly to the lightsheet channel coming in from the right. We then switched on the pivot scan to remove any shadow artefacts and set up a few z-series through the tissue. The image below shows the sort of resolution we was getting off the x5 detection objective using maximum zoom.

Above: Low power reconstruction of neuronal cell bodies in brain tissue. Dataset taken to a depth of 815 microns from the tissue surface  (snapshot of 3D animation sequence).

It took us a fair amount of time to establish a workflow for the correct preparation and presentation of the sample to the lightsheet.  However, once we had established this we were able to get some pretty good datasets and in very good time – the actual imaging part was relatively straightforward. The next step will be to repeat the above using the refractive-index matched X-CLARITY mounting medium,  try out the x20 clearing objective and utilise the multi-position acquisition feature of the software.


Tissue clearing and labelling, I. M. Garay; preparation, presentation and lightsheet imaging of sample, A. J. Hayes; photography, M. Isaacs; text, M. Isaacs and A.J.Hayes.

Further reading: