Chamber Current and Voltage as They Relate to Sample Clearing and Degradation

Perhaps it would be worth a discussion as to the electrical settings involved as they pertain to tissue integrity in the chamber.

My assumption up until this point has been that while the electrical field strength (read, voltage drop over volume) is responsible for the electrophoretic tissue clearing, excessive current will be primarily responsible for any melting or other electrically derived destruction of tissue. Having melted several brains already, I made an attempt today to characterize the amperages involved.

It should be first stated that my chamber is of a different design than Dr. Deisseroth's team. The distance between my electrodes is 7cm, and cross sectional area is 1.425cm^2. This may (read: will) influence the ratios of electric potential drop to amperage. I should, however, experience an increase in resistance in correlation to the increased length between electrodes, and so voltage and amperage will remain directly related by the specific resistivity of solution.

In an attempt to cross check my process with that of the original, I found it necessary to calculate the specific resistivity of solution of the clearing solution. I measured (at 37C), 30V to produce 22 mAmps of current through my chamber.

Using the equation Resistance=length/area*specific resistance of solution, (R=L/A*P), I was able to calculate a specific resistance of the clearing solution to be 277.59 Ohms/cm at 37C.

In an attempt to calculate a "safe" amperage, I moved on to interpolate the amperages achieved by the original paper, using their stated materials and methods. Based on the statement that they were able to run a maximum of 4 setups at 37C and 30V on a 300mAmp-capped power supply (the same that I am using), I was assuming a current in the range of 75Amp per chamber as "safe".

Using the link posted at Clarityresourcecenter detailing ETC chamber construction, I was able to infer that the cross sectional area between Stanford's electrodes is 3.24cm^2, and distance between the electrodes is 1.3cm (the width of the 50ml blue BD bottle cap).

Using these measurements, and my previously calculated specific resistivity value at 37C, and again the formula R=L/A*P, I obtained a resistance of 111.38 Ohms for their chamber. That is to say, with an electrode width of 1.3 cm and area of 3.24cm^2, the resistance of the solution between those electrodes should be 111.38 Ohms.

At 30 volts, the amperage involved in running just one of those chambers is 270mA, already approaching the 300mA cap on the entire power supply. How is it possible to run 4 of these on a 300mA capped PSU?

One thing of interest to me; it strikes me that with constant specific resistivity of solution, and barring any changes in temperature, amperage through the chamber is an accurate measure of voltage drop across a given volume, and so should correlate with the electrophoretic driving force. (and thus clearing speed)

My question to you all is, what sort of amperages are you seeing? What sort of amperages have you been successful with? What damages the tissue? How do you achieve considerable potential drop across multiple chambers without exceeding the amperage caps of your power supplys?

It is possible that my calculation of specific resistivity of solution is off, but it seems unlikely to be wrong by more than 10% or so.


  • I haven't completed my setup yet, so I don't have any amperage data for you yet, but I do have a general question about your setup. Why did you decide to go with the larger ETC chamber? Are you wanting to clear larger tissues or was it for some other reason?
  • To be honest the larger setup was a result of a number of trials during which I had problems waterproofing some very complex chambers. My 4th chamber (which you may have seen in the other thread) is essentially a 3/4" ID clear PVC pipe. I have shorter lengths, but I settled on a 7cm length of pipe with electrodes on either end because that length allowed me to have a viewing window.

    A longer distance between electodes does have the effect of decreasing field intensity, and decreasing rates of clearing - I have compensated by increasing voltage, but do not have the data to establish a safe/working correlation between distance and voltage. Even with 7cm separating the electrodes as opposed to the original 1.3cm, I still melted my last brain while running at 90V (only 3 times higher than "Safe" voltage for the to-spec 1.3cm chambers, although the distance has increased by almost 5.4 times). One would expect that voltage would need to increase in proportion to distance between electrodes, but this does not seem to be the case.

    It has been dawning on me over the past few days that amperage may be a better indicator of clearing capacity, due to its direct relationship to the voltage applied via the specific resistivity of the solution. Perhaps I will make a few runs using constant amperage, and let the voltage change dynamically as the tissue clears.
  • @joelrosiene: In which thread did you post your 4th chamber? We have tried to create slightly more complex, waterproof chambers and have had quite a bit of difficulty doing so.
  • @JosephMarmerstein

    Here are a few pictures of my latest chambers. I am very happy with them - they have given me my best results yet.
  • @joelrosiene
    Thanks for those pictures! Very interesting design. Do you secure the tissue in any way in those chambers (like the cell strainer suggested in the original protocol, for example)?
  • The brains do move, but not by much, they merely bobble around within the room that I give them. I have a mesh cap on each end of the clear section, before the electrode, to prevent the brain from contacting the electrodes and being burned/picking up tarnish. Also, within the chamber, and immediately below the two I/O nozzles, there is a plastic plate which exists to protect the brain from excess turbulence.

    The distance from end to end, (and therefore the distance between electrodes) is 7cm, and I get good clearing ( by running 50V 100mAmps for 2 days.
  • I ran into the same issue yesterday; I set it up to run at constant 40V, but it couldn't even get up that high without an amperage error because my sample was only 118mOhms. I switched over to constant amperage @ ~370mA (the max on my box is 400), and just took whatever voltage I could get without the amperage error. Unfortunately this was only able to get me up to ~10-20 volts. Another observation that I made yesterday was that increasing the flow rate seemed to decrease the resistance in my chamber, leading to a voltage drop; I'm not sure if all builds or samples would end up with the same observation or not, but that was the case for me.
  • @joelrosiene are you just pushing fluid through with a pressure pump? That is a pretty good result after two days, seems like your setups works quite well. How accessible are the electrodes in your chambers? If you are also experiencing the tarnish buildup, do you clean the electrodes before each new sample?
  • Fluid circulator is a cheap Petco fishtank circulator, connected to chamber with PVC tubing. Electrodes in my chambers are just spirals of .13mm Platinum wire.

    The anode sees a bit of black tarnish, as you can see, but nothing major, and I have not found that it interferes with the clearing process, so I don't bother to clean it off.
  • @joelrosiene
    Thanks for answering all my questions. Interesting that you don't see very much deposit on the anode, even after a few days. Our setups accumulate quite a bit of tarnish pretty quickly, so we usually clean them every couple of days.
  • @joelrosiene : that is indeed a pretty good clearing you got, as said by @JosephMarmerstein . Is the sample in a mounting medium, or just straight out of the chamber? Even after a week long clearing, I did not get such a clear result straight out of the chamber. Did you checked if your sample has autofluorescence ?
  • @nicolasrenier : That sample is actually fresh out of the chamber - no RI matching taking place there. I am still looking into what might be an appropriate mounting medium. I'm looking into mixing up some FauxcusClear (clever right?), or perhaps some 80-90% thiodiethanol, as per the SeeDB paper. Also, perhaps ScaleA2.

    That brain is actually a Thy1-GFP. We should have some nice axonal labelling, but per your inquiry, the autofluorescence is causing lots of issues. At this point I cannot even tell if my GFP signal has survived. When I get a mounting media working, I might be able to get a better idea using a confocal scope.
  • stuberlabstuberlab Posts: 9
    @scottyler89 We had the same issue with the current overload. We just threw on a ~500 ohm resistor since we didn't want to continue at such a low voltage. Just make sure the resistor has a high power rating since most are rated at only 1/4 - 1/2 watts (the first one we tried started to fry). It's been working for us so far. I think voltage is more indicative of how fast these brains will clear (and it sounds like high current is turning the brains to goop), so dropping the current with a resistor doesn't seem like it will hurt the process.
  • stuberlabstuberlab Posts: 9
    I take back my suggestion of the resistor. Definitely not worth the gain in voltage. Perhaps clearing is more of a function of power (both voltage and current).
  • @joelrosiene Thanks for posting your chamber design above. It's awesome. We are sick of the little Nalgene cup chambers... everything about them seems to be unstable. Instead, we have been trying to develop a 3D printed design in order to make more custom sizes, but have found it to be a much more challenging issue than expected because detergent has essentially no surface tension and leaks out of anywhere! We are close to a functional 3D printed chamber, but I didn't want to wait any longer to get some samples clearing.

    We were able to find these little threaded PVC pipes in various sizes from McMaster-Carr and are now running a larger version of your design. Took a lot of PTFE tape to get it sealed, but seems to be holding atm. Nice also that we can see inside to monitor clearing!
  • @KatherineHolzem I'm glad to hear that someone else has tried and likes the design! I've found it to be pretty stable. The PTFE tape works well for waterproofing the threading, but be careful about a large surface area of the PTFE being exposed to the clearing solution, because I've seen a waxy residue come off of it which contaminates the sample. I just use a thick strip of it wrapped around the edge of the threads.

    What are your chamber dimensions? How do you think a wider(?) chamber will influence your rates of clearing? How long are your chambers? What voltages are you running at? Are you running at higher voltage due to the greater length of the chambers than in the original design? (As I do?)
  • @joelrosiene... I originally started using chambers of the same length as yours (7 cm), and I have gone only as high as 50V. Things seem to heat up too much (for the tape and tube to withstand) if I go any higher than that. I had to switch from PTFE tape to silicone tape, which seemed to hold up a bit better... that detergent seems to find it's way through anything.

    I have made some of my own modifications to your chamber design at this point though. I found these Fernco Qwik Cap things at Home Depot, and I think they are totally awesome for sealing the ends. Here is a picture of the design:

    Basically, if I was actually able to get the threaded cap to seal, I had a very difficult time getting it off. These caps are flexible PVC with a cuff for tightening. Forgive the ugly white PVC pipe at the moment. I just wanted to test the caps in principle. I am actually thinking of switching to a clear polycarbonate tube now, which will have much greater temperature resistance. Not having threading also means I have the liberty to make the chambers shorter. I have also experienced a much smaller amperage with the larger spacing between electrodes, and I am guessing this is because the resistance is greater.

    Not completely sure how the width of the chamber will impact the process, but for a given length, the resistance is less for a larger diameter. Also, the nice thing about the pipe design is the majority of the current flow should be through the pipe (and sample).
  • @KatherineHolzem looking good! I'll just add a few comments towards the challenges you brought up.

    I addressed the heating issue (which occurs at the ends of the chamber) by manipulating the flow of the solution. Inside each of my chambers there is glued in place a plastic breakwater of sorts which both protects the sample from turbulence and ensures that solution flow through the chambers actually occurs, rather than simply an area of high flow between the in/out ports and the rest of the chamber being relatively stagnant.

    As for the PTFE tape - I agree that the detergent is very good at finding its way around it. I use a pretty thick, but very skinny, wrapping of that about just the ends of the threads. Good suggestion with the silicone - I will have to acquire some.

    I tend to use vice-grips to remove the caps after a few days of clearing :)

    I actually began this thread because I wanted to better understand the relationship between Voltage, current, and the rates of clearing that we see. My initial impression was that because the clearing is electrophoretic, we would be interested in maximizing voltage drop across the tissue (placing the electrodes as close to the tissue as possible) in order to best push the detergent micelles through the sample. I am not 100% on this.

    I think we can agree that increasing the width of the chamber, even though it does provide a higher electric current through the chamber, is probably not desirable. I would guess that the optimal chamber width closely matches that of the sample, without occluding solution flow on either side of the construct. Also, one should account for the expansion of the sample during the process.
  • ButchButch Posts: 1
    I also started out using a similar "reticulation tubing" design, almost identical to those used above... but found leakage and opening/closing too tricky and messy. I've now gone back to the Nalgene chamber design, but with a fundamental change. I'm using those 'squirty' nalgene bottles, cutting the inner tubing, then attaching half of a 10ml syringe which houses the tissue to be cleared... I make sure the hose connections aren't near the SDS fluid, and thus I don't need glue (the reticulation connectors screw into the bottle nicely). The anode Pt wire sits in the bulk of the fluid, and the cathode in the neck of the bottle tube.. This way all of the fluid and current flows through the 10ml syringe and brain (high density). Theoretically, this design should also allow the dissolved lipids to float to the top of the fluid in the bottle, but the 10ml syringe only draws 'fresh' SDS from the bottom of the bottle (not that I've ever seen floating fats after several days). A little gauze in the 10ml syringe stops the brain getting sucked up and blocking flow. Easy to unscrew and change brains, and doesn't leak.
  • I was wondering what the most up to date ETC chamber design is, whether it be 3D printed or DIY. I have been working on a design lately and just want to know if im fruitlessly trying to reinvent the wheel due to someone already having a new really good ETC chamber design. Pictures?
  • @Solder I must say that I am quite attached to my chamber design (seen several posts above), and have been using it successfully for some recent runs. @KatherineHolzem and @Butch have both made some significant improvements in terms of preventing leaks.

    The 3D printed chamber that I tried out had a little bit of a leak, which I might be able to fix with a better O-ring.

    I think you will find that the biggest challenge in building your own chamber is going to be preventing leaks while at the same time having simple and easy electrode access.
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