I've been thinking about trying the CLARITY method myself since Deisseroth's paper came out. I'd love to find out how well it works and some of the pitfalls you encounter Will. Thanks for taking the initiative and sharing your experience with the neuroscience community. If you'd like to collaborate or work on it together, I'd be happy to do so, seeing that I'm not too far from where you are.

By the way, do you think a CLARITY kit would be commercially available soon, just like Scale solution? He has got to be in the process of commercializing it either himself or in partnership with a company like Olympus or Invitrogen. However the price of the kit would probably be much higher than the sum of all parts on Deisseroth's materials list.

In the spirit of sharing, I've been using Scale solution for a year now clearing brains that had been IUEd. Scale works great and is, like you said, economical. Don't buy it from Olympus, since they charge something like $300+ per liter, when you can make it yourself at a fraction of the cost. Just remember to use DI water, otherwise it may not work. Scale works really well on thick brain slices, such as those used for ephys. We can image through and through a 4oo um slice on a confocal LSM with ease. On whole brains it works fairly well but takes much longer. We tend to hemisect the brain before clearing. This not only helps with clearing efficiency, it gives you more flat surfaces on which to orient your brain during imaging. Here are the down sides to Scale. 1) On a whole or hemisected brain, optical sectioning of deep brain structures is still not ideal. Resolving fine structures such as axons and fine dendrites is still difficult, presumably due to diffraction limits. Even fluorescent somata are sometimes difficult to resolve cleanly in whole brains cleared for 2 weeks, if they are too densely packed such as in the rostral migratory stream. 2) The cleared tissue swells approximately 20-30% so everything looks a little bigger, making absolute measurements of cell parameters impossible. Relative comparison between tissues cleared in Scale is okay. 3) While gray matter clears beautifully, white matter such as the corpus callosum is hard to clear. Tracking a single labeled axon across the corpus callosum in a thick slice cleared in Scale is not feasible (at least in our hands). 4) Once cleared in Scale, the tissue needs to stay in Scale or the opacity will revert, which means submerging water immersion objectives in high concentration of urea in order to image in magnifications larger than 20x. We don't know if this shortens the life of the water immersion objectives.

Dear all ( reading this post - and from the number of views within the last 15 mins I can see that this question might be of interest to many who are not knowing about that method-),

just to cite the reference for the mentioned new "CLARITY" procedure (since I did not find any result googling the term < Clarity > or < Clarity clearing >:

cf.

Chung, K., J. Wallace, S.-Y. 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. 2013. Structural and molecular interrogation of intact biological systems.

Nature advance online publication(April).

[source: http://www.biotechniques.com/news/I-Can-See-Clearly-Now-Through-a-Mouse-Brain/biotechniques-342028.html?utm_source=BioTechniques+Newsletters+%26+e-Alerts&utm_campaign=4b6f04267a-daily&utm] .

Citation: < Mouse brains have been made transparent before, most notably by using the SCALE [cf. http://www.biotechniques.com/news/biotechniquesNews/biotechniques-325032.html?utm_source=BioTechniques+Newsletters+%26+e-Alerts&utm_campaign=db25ae3943-Methods+Newsletter&utm] method developed in 2011 by Japanese researchers. But CLARITY is different, said Deisseroth, because it allows “not just transparency but labeling within the transparent organ in a very flexible, versatile way. You can use any molecular label to paint different neurons different colors and see which are excitatory and which are inhibitory. CLARITY can add a lot of information that was not available before.” >

Best regards

I haven't try these techniques, but the wide coverage of the Deisseroth CLARITY paper might overshadow previous reports who did pretty much the same thing, in 2007! One thing for sure is that Deisseroth protocols seems details and an improvement on previous techniques.

Nat Methods. 2007 Apr;4(4):331-6. Ultramicroscopy: three-dimensional visualization of neuronal networks in the whole mouse brain. Dodt HU, Leischner U, Schierloh A, Jährling N, Mauch CP, Deininger K, Deussing JM, Eder M, Zieglgänsberger W, Becker K. http://www.nature.com/nmeth/journal/v4/n4/pdf/nmeth1036.pdf

It's too bad that the Scale technique did not come to my attention earlier, because it should also be good for our purposes.

It seems that the main advantage for CLARITY is that, according to Chung et al., it only results in ~8% protein loss, versus ~41% for Scale. I wonder if this will make a difference for us; we are just hoping to visualize IUE-transfected GFP neurons. Any IHC benefit is just a perk that we may or may not take advantage of.

The main advantage for Scale seems to be price and simplicity (no electrophoretic tissue clearing is necessary). We're going to compare both techniques; I'll try to report back in a few months.

Here's the citation for the Scale technique: Hama H, Kurokawa H, Kawano H, Ando R, Shimogori T, Noda H, Fukami K, Sakaue-Sawano A, Miyawaki A. 2011. Scale: a chemical approach for fluorescence imaging and reconstruction of transparent mouse brain. Nat Neurosci. 14:1481-8. http://www.nature.com/neuro/journal/v14/n11/full/nn.2928.html

I should also add that the full CLARITY protocol can be found at http://clarityresourcecenter.org/

I've been thinking about trying the CLARITY method myself since Deisseroth's paper came out. I'd love to find out how well it works and some of the pitfalls you encounter Will. Thanks for taking the initiative and sharing your experience with the neuroscience community. If you'd like to collaborate or work on it together, I'd be happy to do so, seeing that I'm not too far from where you are.

By the way, do you think a CLARITY kit would be commercially available soon, just like Scale solution? He has got to be in the process of commercializing it either himself or in partnership with a company like Olympus or Invitrogen. However the price of the kit would probably be much higher than the sum of all parts on Deisseroth's materials list.

In the spirit of sharing, I've been using Scale solution for a year now clearing brains that had been IUEd. Scale works great and is, like you said, economical. Don't buy it from Olympus, since they charge something like $300+ per liter, when you can make it yourself at a fraction of the cost. Just remember to use DI water, otherwise it may not work. Scale works really well on thick brain slices, such as those used for ephys. We can image through and through a 4oo um slice on a confocal LSM with ease. On whole brains it works fairly well but takes much longer. We tend to hemisect the brain before clearing. This not only helps with clearing efficiency, it gives you more flat surfaces on which to orient your brain during imaging. Here are the down sides to Scale. 1) On a whole or hemisected brain, optical sectioning of deep brain structures is still not ideal. Resolving fine structures such as axons and fine dendrites is still difficult, presumably due to diffraction limits. Even fluorescent somata are sometimes difficult to resolve cleanly in whole brains cleared for 2 weeks, if they are too densely packed such as in the rostral migratory stream. 2) The cleared tissue swells approximately 20-30% so everything looks a little bigger, making absolute measurements of cell parameters impossible. Relative comparison between tissues cleared in Scale is okay. 3) While gray matter clears beautifully, white matter such as the corpus callosum is hard to clear. Tracking a single labeled axon across the corpus callosum in a thick slice cleared in Scale is not feasible (at least in our hands). 4) Once cleared in Scale, the tissue needs to stay in Scale or the opacity will revert, which means submerging water immersion objectives in high concentration of urea in order to image in magnifications larger than 20x. We don't know if this shortens the life of the water immersion objectives.

@ William Adler: Dear William, would you mind to check the reference URL for the CLARITY protocols again since I got an error message: ....server does not have a DNS entry....but it may well be that it is a probem on my (our server) side... Thanks also (also @ Lawrence Hsieh) for your posting essential and acknowledged information on those two marvellous techniques ( I unfortunately will not be able to work with these despite having great interest! having done semithin section serial reconstruction of magnocellular nuclei in rat hypothalamus for my PhD thesis)... Best wishes and good luck, I shall follow this post eagerly...

Wolfgang: Sorry, I pasted the wrong link into my comment. The link won't work if it contains "www." Go here: http://clarityresourcecenter.org/

Lawrence: Many thanks for the tips on Scale. We have also been thinking that hemisecting is probably a good idea, so I'm glad to hear that you do it.

It's unfortunate that you could not trace axons across the corpus callosum; that's something we will want to do. It sounds like Scale is worth trying out, but I'll be interested to see whether CLARITY is a substantial improvement in image, ahem, clarity.

1) Can you resolve axons well with Scale when using a 400 µm slice?

2) Why do you need to immerse the objective in Scale? Would it not be possible to put together a glass imaging chamber, as in the CLARITY paper? That way you could immerse the objective in water, rather than Scale.

3) Do you use mice or rats? We will be using rat brains. I wonder if we will have to alter the Scale or CLARITY protocol (incubation times, etc).

By the way, after ordering the supplies on the CLARITY protocol, the total startup cost was only about $3k. We ordered a less expensive Lauda-Brinkmann immersion circulator than the one recommended, and we already have a power supply and many chemicals, but this should be a pretty affordable project, barring any unforeseen issues...

I'll try to post updates on this thread, and yes, let's consider the possibility of some kind of collaboration, depending on how this goes!

Dear William, thank you for the new ( correct ) link to CLARITY....(:-))

best wishes and regards, Wolfgang

Dear All, I am also very interested in trying the CLARITY method myself. I have tried already the Scale and the BABB-based method and both have problems such as a very long time necessary for clearing to take place (in case of the Scale) and the fluorescence bleeching and light scattering (in case of the BABB). I must say though, that I study neuronal circuits in adult rat brains and spinal cords and adult tissue definitely presents a bigger challenge when it comes to clearing and imaging. I sure hope the CLARITY will resolve problems of the previous two methods. Best regards,

To answer your questions Will,

1) We can certainly resolve axons that come out of electroporated cells, watch them cross the corpus callosum and emerge in the contralateral cortex in 400 um slices, but to follow a single axon from one pyramidal neuron across the corpus callosum and its collateralization on the contralateral side is currently not doable in our hands, even in cleared slices.

2) Immersion of the objective in Scale is not necessary if you are using an air objective. However resolution, or numeric aperture, and working distances are limited in air objectives. I've not tried putting cleared tissue in FocusClear, as described in Deisseroth's protocol, but in other liquid media other than Scale. The outcome has been invariably the same, and that is the tissue reverts back to its opaque state to varying degrees.

3) We've cleared mouse brains from new born to adults. Scale works equally well across the board and is easily scalable temporally. The advantage of Scale is that you can see the tissue clear before your eyes, which allows you to empirically determine when your tissue is suitable for imaging. I don't think you'll have a problem clearing rat brains, especially if you hemisect before clearing.

Clarity is an impressive technique with only a claimed 8 % protein loss - would be good to see the results from other labs

Mario, are you not able to order it through Wako Chemical?

http://www.wako-chem.co.jp/specialty/waterazo/VA-044.htm

We're anxious to try our automated neuron reconstruction software, AutoNeuron, on large tissue specimens like what could be generated with the clarity and scale protocols. We've done it successfully on specimens over 1 mm thick - if you have image stacks with labeled cells/fibers in thick tissue that you'd be willing to let us play with, let me know!

If anyone else in the US is having trouble buying FocusClear from CelExplorer labs (or wants to avoid the shipping charge), you can buy it from their US distributor, Cedarlane Labs.

https://www.cedarlanelabs.com/Products/Search?lob=All+Products&searchby=Description&supplier=CELLEXP&text=&x=51&y=3

I'm trying CLARITY in our lab right now but am having trouble getting the hydrogel to polymerize. Has anyone gotten past this step?

We just did that last week, and had success. Are you making sure to degas your tubes with nitrogen? We were skeptical, but it really is crucial to flush out the oxygen.

William Adler - Thanks for your response. I degassed 3x with nitrogen in a small vacuum desiccator and then put the desiccator under vacuum again and put the whole thing in a 37 degree oven. I was worried that opening the desiccator lid to close the tubes would cause too much oxygen to rush in, so I just kept it under vaccuum throughout the polymerization. Now that I think about it, maybe that was not such a good plan. I was also unclear as to whether the sample should be rotated or left still during polymerization. In the protocol, it almost sounds like it should be on a rotator in a 37 degree water bath, which was confusing to me.

Hey Will-

I'm working on getting the Clarity protocol up and running using mouse brains and for the first time this week will be trying to clear using our homeade electrphoresis chamber. I'm interested how you fabricated your circulation system and if you've successfully cleared a brain yet? Are you starting off with whole brains or thick slices?

I agree; I'm not sure why they mention a rotator on the protocol. We just submerged the tube in a water bath and left it still.

We were also worried about the level of oxygen exposure if we followed protocol and opened the desiccation chamber to rapidly close the tubes. It definitely felt a little sketchy, but it worked. And our non-degassed control tubes did not polymerize.

I'm not sure why putting the desiccator in the oven didn't work, but maybe try the method in the protocol.

Also, we are currently clearing our first brain in the ETC chamber. I'll report back later in the week on how it goes... fingers crossed.

Nick: We built our ETC apparatus almost entirely from the part list on the protocol, except with a 125 mL Nalgene Straight-Sided Wide-Mouth Jar instead of a 60 mL version. We also bought a Lauda-Brinkmann A6 circulator, which lacks many of the features of the RE 415, such as cooling.

We just tried to clear our first whole rat brain. We ran it at 50 V / 50°C. After 24 hours it had swelled grossly, filling up the whole cell strainer, and being unrecognizable as a brain. It's also very soft and unsturdy.

For our next run, we are going to try one cerebral hemisphere instead of a whole brain. We might also try lowering the voltage and temperature; what we ran was close to the maximums recommended in the protocol.

Hey Will-

Thanks for the hardware info. We're starting on the lower end (20V @ 37C) and running just during the workday and not overnight until we gain some confidence in the system. Going to be a slow process with low voltage/temp and running only 8hrs/day. Hopefully we can bump that up to running 24hrs shortly. I'll keep you updated, and will be interested to hear how your second round went. I think lowering both the temp/voltage would be a good thing for you to try.

Elizabeth Kritzer - how long did it take for your hydrogel to polymerise once you had it in the water bath? I am having issues with getting this part to happen, even after degassing the sample to the letter.

Thanks.

Hi

I have polymerized my Brains without Problem. Just follow the protocol, giving extra care to keeping all cold all the time.

The first Gel took 4 hours to become really solid. After ca. 3h was kind of solid but still little viscous. Leaving the Gel polymerize o/n showed no changed comapred to 4h for me

Cheers

Fabio,

Have you successfully cleared a brain after embedding?

Hi William,

Sadly im still constructing my ETC chamber. Macmaster Carr doesnt ship to Germany.

So I had to invest some time looking for components here. I hope in a couple of weeks Im ready to go.

In the mean time I will try some other ideas after embedding.

Hope to hear some good news from you after you cleared your brain.

Adam - I'm still not able to get my gel to polymerize, even though I am doing exactly what the protocol says. I suspect my degassing setup is not working. I might try bubbling with N2 before degassing, to see if that helps. Can I ask what kind of degassing chambers/desiccators others are using?

Additionally, I'm making my own PFA fresh from powder instead of buying the premade stuff listed in the protocol. It seems unlikely that this could be what's causing the problem, but can anyone confirm that using fresh-from-powder PFA works in this hydrogel?

Elizabeth,

We are using a Bel-Art desiccator: http://www.belart.com/shop/420270000-space-saver-vacuum-desiccator-230mm-clear-p-420270000.html?y=12&x=66

We too are making our 16% PFA from powder, pH adjusted to 7.4. We are using the other ingredients from the vendors listed in the protocol (Gibco/Invitrogen, Wako, Bio-Rad).

Can you confirm that you are getting a strong vacuum seal? With our desiccator, the lid can't be removed under vacuum.

We are on day three of clearing our second brain at a gentler temperature/voltage. It's going slow, but it's not getting soft and swollen like our last one, thankfully. We've been taking it out to check on it every day, and I'm just now realizing that I haven't been maintaining the orientation of the brain when I put it back in. Would you all think that it's important to keep the brain's orientation consistent, with respect to the electrodes, for the ETC to work?

Nick, how is clearing going for you?

We are currently clearing a mouse hemisphere at 20V and 35C. I had trouble polymerizing a batch of hydrogel solution + saponin but repeated the N2/vacuum/N2 process and it worked like a charm. I think you really need to make sure you flush the desiccator well with N2 and ensure you have a good seal for the vacuum - I discovered that the lid had not formed a tight seal my initial attempt so it wasn't pulling a very good vac.

How do you all plan on imaging such a large piece of tissue? We have an inverted SP5 so using a dipping lens from the top is more troublesome (requires objective inverter). Also the paper describes their technique but I'm still a little fuzzy on how they constructed their imaging chamber (using Blutack). They also mentioned imaging one half, inverting, and imaging the second half. I know you can merge image stacks using XuvTools - does anyone know exactly how they went about merging this huge data set? Even using a resonant scanning confocal, to collect and stitch together such a large volume will likely require hours of scope time.

For interest, here is our first (moderately) cleared rat brain, in a whole-brain imaging chamber made of a slide, a Willco dish, Blu-Tack, Kwik-Sil, and 80% glycerol (we're saving the FocusClear for a brain that might actually be clear enough to image!). We had to scratch the slide in order for the Kwik-Sil to have a rough surface to grab onto. It appears to be watertight at this point...

Side view.

David,

Although we have built an imaging chamber, we actually don't know how we will be collecting and combining our data. Once we get a good and clear brain, we plan to work with our confocal imaging center to figure this one out; they seem to think that we have the equipment, but no one has done this kind of imaging.

William-

Our brain is actually looking quite similar in coloration to the images you posted above. Starting to clear a bit, but far from transparent (what looks like transparent tissue is actually hydrogel remaining from embedding step). We're going to keep running it in the ETC chamber and see if it continues to clear. So far its only been running for a total of approx. 40 hours (published protocol said up to a week running at 24hrs/day) at low voltage/temp.

We're working on getting more ETC chambers set up so that we can run multiple brains in parellel.

William -

Thanks for the pictures. That gives me a better idea of what they may have been describing.

Has anyone tried clearing tissue other than brain yet? We only have one ETC chamber at the moment but plan on trying kidney, liver, and pancreas.

Thanks for the pictures, guys. That helps me understand how to set up the slide once we get something cleared.

Will - I would imagine that you'd want to put the brain back in the same orientation each time. As I understand it, the lipids will migrate from anode to cathode, eventually leaving the tissue on the cathode side. If you run it for a while, then flip the brain around, the lipids might move back towards the center of the tissue (since now the cathode is on the other side). I probably didn't explain that very clearly, but basically I would leave the brain undisturbed if possible to maintain consistent migration. Also, thanks for the desiccator info. Looks like you're doing the exact same thing I'm doing, with the same equipment...so I must be technically failing somewhere; time to keep troubleshooting.

David - Our lab is mainly concerned with the spinal cord, so that will be the first tissue I try to clear, rather than brain. I will certainly post about my results once I get things working here.

Nick,

That's about what our current attempt was looking like when we checked on it this morning (after running it for 3 days at 15-25V/37C). Hopefully we are on our way! Are you making sure to put the brain back in the same orientation? I think Elizabeth's advice is probably right.

Elizabeth, did you cool down all of your ingredients before you started mixing the hydrogel solution?

Will

Will - maybe that was the problem. It's possible that my stirplate was residually warm from some other solutions I prepared that day.

I remade the hydrogel today, taking extra care to keep everything cold and holding vacuum for 30 min. straight when degassing. 2.5 hours later, I have almost complete polymerization. I did bubble a tube with N2 for 30 min prior to degassing as well, but it didn't seem to make a huge difference; the bubbled tube seems to have polymerized a little faster, but I bet the non-bubbled tube will catch up in another hour or so. I am guessing that in my first few attempts I did not get a sufficient vacuum after only 10-15 min. Even though I did several vacuum/purge cycles in a row, maybe it required 30 min of constant vacuum to pull all the O2 out. I also did not freeze/thaw my hydrogel before polymerizing. I will have to thaw some and make sure it still works before I try perfusing an animal with it.

Thanks to everyone who helped with info and suggestions.

Bertrand,

The design in the protocol includes a circulator that has a fast and uncontrollable flow rate. But it also includes a stopcock in the circuit that can be used to adjust the flow.

Will

Yes you can adjust the flow using a stopcock, we even have 2 in our setup. The length of the tube can also affect flow. The important thing with the flow is to prevent bubble formation

Thanks for starting this discussion. I have gotten a CLARITY setup together in my lab and have been able to clear a mouse heart at this point. To answer Elizabeth's question from above, I do think that making the 16% PFA is the problem with your polymerization. I don't know why this is, but I have made several batches of hydrogel solution at this point, and the only batch that didn't polymerize (or, I should say, just polymerized very, very slowly) was the batch with homemade PFA. Every time I have used the pre-made stuff from EMS, I have had no problems. My assumption has been trace remaining paraformaldehyde polymers may impact acrylamide-bis polymerization, but I don't know this to be the case... just know that I am ordering the EMS 16% PFA from now on because it's not worth potentially ruining tissue samples for how cheap this solution is to buy.

The main issues that I have come across, and I would like to know if others have experienced the same, are with the electrophoretic clearing. First of all, fabricating a reliable water tight chamber has been more difficult than expected, and this is still a work in progress. For the Nalgene bottle chambers, I would not recommend the hard epoxy that the Deisseroth lab uses. It cracks too easily, and then you just have a leaky chamber. Casting Sylgard on the inside has worked the best in my limited experience. Ultimately, some labmates and I are trying to come up with better 3D printed designs for these chambers. Secondly, in running the electrophoresis continuously, I am getting a black deposit on the anode (and, more problematically, the tissue). It does wash off, but it accumulates quickly enough that things need to be cleaned pretty much daily. Also, the 0.5mm platinum electrode that they recommend seems prohibitively expensive. I have used wire as thin as 28 gauge successfully, but am now going to test some platinum-iridium alloy.

Let me know if you have thoughts or suggestions on these issues. I would say that the protocol that the Deisseroth lab put up was pretty awesomely detailed, but still, getting a reliable system running, has been fairly challenging... involving a lot of chamber-sitting and cleaning up of SDS clearing buffer.

I can try to jump in the conversation. I tried to read through the whole thread, but it is getting very long now!

I've been trying the Clarity protocol since the day it was published, and here are some thoughts. Tell me what you think :

- The brains seem to be always yellowish after the ETC clearing. I don't know if it is supposed to be like that, but everyone I talked to has the same problem.

- The brain after the ETC clearing expands a bit, but is not really transparent. It is slightly light permissive, but not "see through"

- It is very counter-intuitive, but the brain should move freely in the chamber. Do not try to secure the brain in the same position. It should be tumbling. Don't ask me why! Prefer long runs (4 days to 1 week) at low voltage and low temp (30V, 37°c). Never interrupt a run to look at your brain.

- Black deposit occurs on one electrode, even with platinum. There doesn't seem to be a way to prevent this.

- The only way to get a completely clear brain seems to be though the use of a clever mounting medium (what they call "index matching" in the paper. The focusclear seems to be the best way to get to this point. ETC clearing will not give you full transparency. Glycerol does NOT work. ScaleA2 is great as a mounting reagent, but it defeats the purpose of "Clarity" for many reasons. A good way would be to develop a mounting reagent with high refractive index (high molarity). Using DMSO and diatrizoic acid could be a good start.

I hope this helps. Let us know if someone gets to the end of this protocol successfully!

Also, confocal imaging will be impractical for large fields of views. The logical thing to do would be to try light sheet illumination (SPI microscopy)

Katherine - I also had issues with the watertight chamber, but ultimately found that Radio Shack's Quick Setting Epoxy works really well. I put a thick coat on the inside and outside of the inlet/outlet ports, and just the outside of the electrode ports to hold the wires in place and seal. Then I let it cure overnight and no problems with leaking despite running my setup for several weeks.

I did my first trial run with a spinal cord this week, but had to abort the clearing early due to cracked tubing. We are using a peristaltic pump to circulate, so I imagine this will not be an issue for anyone using an actual water bath circulator as in the protocol. Anyway, I noticed some clearing and yellowing of the tissue, as Nicolas stated. It's good to know not to expect full transparency after only ETC with glycerol...guess we will have to invest in some FocusClear.

Nicolas - I'm intrigued by your DMSO + diatrizoic acid clearing method. Can you provide the recipe you use? I'd like to give it a shot.

@Elizabeth and Katherine : I agree with Elizabeth that the epoxy recommended in the protocol is not good for the Nalgene plastic surfaces of the chamber. I also used an off-the-shelf epoxy from 3M that is designed for plastics and cures much faster. to completely seal the system, I use teflon tape at every joint to be sure there are no leaks.

@David : The mounting reagent is still a work in progress, unfortunately. If you're in a hurry, you should rather use Focusclear. Because I didn't want to spend about 100$ per brain of mounting reagent, I am trying, with the help of friends at Princeton, to find a home-made replacement.

A good place to start is to look at the Focusclear patent. So here is what I suggest for a start, but you should know that it is not yet optimized. Use DMSO as a solvent, and saturate with an equimolar mix of diatrizoic acid and sodium diatrizoate. You can start at 20% w/v (10% each). To dissolve, use a sonication bath or warm gently at 50°c.

Then, in the patent, they mention the presence of meglumine (N-methly-glucosamine). Diatrizoic acid is used in medical imaging and administered to patients with an equimolar ratio of meglumine. SO you can add meglumine accordingly to the amount of diatrizoic/diatrizoate molecules you have, but you can try without meglumine.

Add 0.1% of tween20, and if you're fancy, you can try to add NADPH and EDTA (I haven't tried yet). millimolar end concentrations should be a good place to start.

I'm sorry it is not a precise recipe, but I am still trying to figure out the best mix to have a great clarification. Let me know if you have more ideas, or what do you find if you try this recipe.

@Nicolas- Curious as to why you say you should never interrupt a run to look at your brain?

@Nicolas - Can you elaborate on the difference in appearance between the secured and free-tumbling brains you tried? I was planning to anchor my spinal cord such that it stays parallel to the electrodes, but I am curious as to your findings with the secured brain.

@Nick : I agree with you, it doesn't make much sense. Those two remarks (the brain should be tumbling, and don't look until it's done) are from the paper's first author who was helping my colleagues to set up the system in their lab. They passed on these informations to me. So it's kind of a second hand information. I think it is not a game changer : I did interrupt my runs at the beginning to monitor the process, and it seems ok.

However, it did seem to work better if the brain can move a little bit freely, and if you don't interrupt. I feel that it is important that the brain receives a constant supply of fresh buffer. So the flow would be important, and I am now using the maximum flow given by the circulating water bath (about 20 liters/min). I considered peristaltic pumps at the beginning, but the flow can not be that high with those. During the runs, the electrodes produce a lot of heat, that has to be quickly dissipated by a rapid supply of fresh buffer.

I should point out that I am still discovering the protocol like everyone here, so I would be pleased to be challenged on my remarks if someone as a better experience with the system!

@Katherine: I am working on clearing a mouse heart and am wondering the conditions you placed yours at for the electrophoresis and how long it took you to clear it. Also do you have any pictures you could share of the ending result. I am still having trouble creating a water tight system so I have not been able to run it continuously very long, but am going to try some of the suggestions listed above.

@Elizabeth : In my experience, it really doesn't hurt if the tissue is freely moving. You have to be careful that the tissue never come too close to the electrodes though. I burned a sample that once came too close. I use a mesh pad to make a sort of isolation between the electrodes and the sample (see the picture). You can buy it on amazon.

obviously, securing the sample doesn't prevent the electrophoresis process from occurring. But the surprising thing is that not securing the sample seems to be at least as efficient. Maybe it is just luck, but I always had bubbles trapped between the cortex and thalamus before, and it didn't happened as much with freely moving samples. The rationale is maybe that the lipids micelles can then exit the tissue from all sides without having to go through the whole brain?

Just to let everybody here know that a forum has been opened on the official Clarity website, so it would be the best place to ask all those questions and continue the discussion!

http://forum.claritytechniques.org/

@Amanda. Here is a mouse heart, and it has taken about 10-14 days at 30V to get them pretty clear. If anyone else is trying heart tissue (or anything other than neural tissue), I would be interested to hear how imaging has gone for you. I have noticed a lot of autofluorescence in the red and green wavelength ranges. I will be testing blue and far-red dyes next week to see if I can just avoid the heavy autofluorescence region, as I don't know if it can be quenched or removed.

@Nicolas, thanks for sharing the link to the clarity forum site, and for your suggestions on a FocusClear alternative.

@Katherine : your heart looks good, I think! In the right mounting medium, you might have a much better transparency. For the brain tissue, I also have a very strong autofluorescence in the green and red. Blue is somewhat better. I'll try to see if changing the formulation of my experimental homebrew focusclear can improve on that.

I will be going to Stanford to see the process in person and discuss with the Deisseroth lab. Send me a message if you have any questions you would like me to ask them.

@Nicolas. Yeah, the heart did actually get a bit clearer than this with just the ETC process, but it sort of broke in half, so this is the best whole picture I have currently. I have not taken a post-FocusClear picture yet, but it does seem a bit better after incubating in it for a few days.

@Bertrand Thanks for pointing that out. I made some progress recently with the fake focusclear, but I'll try this solution on ETC treated samples.

Also, reading this paper reminded me of this idea that the yellow tint taken by the tissue during the electrophoresis could be due to the Maillard reaction, and could be the cause of the very high autofluorescence. Maybe adding a reducing agent in the clearing solution could help?

Nice image of your preparation Katherine - very encouraging and clear

We also have been performing CLARITY on mouse kidney with pretty good results. Using 2-photon microscopy on top of CLARITY, we are getting good resolution and depth of imaging. I'll post pictures once we transfer them off of the workstation computer.

In addition, the Deisseroth lab recently put out a pretty thorough methods paper re: CLARITY in Nature Protocol. There they describe the COLM approach...anyone out there with experience with this approach to imaging?

http://www.nature.com/nprot/journal/v9/n7/full/nprot.2014.123.html

Most clarification methods such as Clarity, seeDB, Scale, are preceeded by an incubation period in 4% PFA of the sample before it is clarified. The length of incubation is typically 12h - can longer incubation times impact the clarification so it fails?

 What is the best purchase strategy for the whole clarity setup in Europe? Best Petr Zach

Hi all components/reactives can be purchased in Europe.

I reccomend you to check the CLARITY forum and wiki

http://clarityresourcecenter.org/

 In case anyone still stumbles on this (like I did), Logos Biosytems has a complete system for CLARITY that cuts the processing time to < 3 days. I haven't used it myself, but the pics on their website look pretty impressive.

http://logosbio.com/x_clarity/x_clarity/features.php