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.

AJH.

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

• Olympus BX61 fully motorised system: Upgraded with new PC (Win 10) and 4K widescreen monitor, Hamamatsu Spark CMOS camera, CellSens Dimensions software and new CoolLED pE300 white LED illumination source.

• Olympus BX40 system: Upgraded with new PC (Win 10) and widescreen HD monitor, Hamamatsu Spark CMOS camera, CellSens Standard software.

• Olympus BX41 Surveyor Slide scanner system: Upgraded with new PC workstation (Win 7) and widescreen HD monitor.

• Olympus BX50 system: Upgraded with new widescreen HD monitor.

• Olympus SZX12 stereo zoom system: Upgraded with new PC (Win 7) and widescreen HD monitor, Orca ER2 firewire camera and HCI image software.

• Meiji macro system: Upgraded with new PC (Win 10) and widescreen HD monitor.

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.

AJH

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.

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.

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.

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)!

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.

AJH

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).

Contributors

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.

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.

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.

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).

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).

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).

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!

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

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

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)   

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).

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).

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).

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).

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.

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

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).

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).

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.

Contributions:

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:

• Zeiss: Lightsheet Z.1 Product Information