Tag: lab

Practical uses of 3D printing in an electrophysiology lab

I have mentioned before about my 3D printer, and how useful I have found it. Today, I’ll explain some of the practical uses I’ve found for 3D printing for electrophysiology. The point is that electrophysiology equipment is both extortionately expensive and annoyingly non-compatible. So, it is often quicker, cheaper and easier to design and print a “thing” than try to buy something to fit your particular need.

Build-a-bath workshop

A year or two ago, I was setting up our third (at the time unused) rig for calcium imaging. We had various bits of baths, but no complete set. And it would cost a (relatively) large sum of money to buy a replacement. I found that what I was missing was the “bath” bit (I had the holder). So I measured up one of our existing ones and designed a reasonable copy in Autocad:

3D design for an electrophysiology bath insert.

My printer was able to make it with a very smooth base, which is the crucial aspect for obtaining a watertight seal. I installed it on the calcium imaging rig, it worked well, and is still in use there to this day. Oh, and for anyone who’s interested, I have made the 3D design available on Thingiverse.

Moving an LED

My loyal readers will know that it was around this time that the light source for the calcium rig died. A replacement LED source would cost anywhere from £3k to £15k, depending on how many colours I wanted access to. However, we had animals ready for experiments at the time, and even if we had the money, it could take weeks for new kit to arrive.

Luckily, we had a blue LED of the correct wavelength attached to one other rigs, that was no longer needed there. I had purchased it to do optogenetic stimulation, but we had switched opto’s to the third rig.

Anyway, this seemed like an easy fix, just swap it over. But of course, it’s never that easy. Because, I wanted to move the LED from an Olympus microscope to a Zeiss. And the manufacturers do not make it easy on the consumer by having common fittings.

So, I measured up the fitting on the LED and the back of the Ziess fluorescence port. I then designed a 2-part “sleeve” that would modify the Zeiss port to resemble the back of an Olympus:

3D design for an Olympus-to-Zeiss microscope fluorescence adapter.

I used cable ties to hold it on tight to the Zeiss fluorescence port. The benefit of cable ties over something more permanent like glue is that they can just be cut off if/when the LED wants changing. The LED now fitted snugly onto the back of the microscope, and, after some fiddling with the data and control connections, was now fully functional for calcium imaging.

A “lab things” service

The main point I want readers to take away from this post is the usefulness of 3D printing for electrophysiology labs. I would strongly recommend anyone reading this who performs a practical skill in the lab like electrophysiology to consider investing in a 3D printer. They are actually quite cheap nowadays (mine was about £250 a few years ago), and I’m sure they’ll save you a lot of time and money in the long run.

In fact, the biggest investment to 3D printing things yourself is the time it takes to learn 3D CAD software and optimising the 3D print process itself. So, if you want something custom making, but would prefer not to have to figure it out yourself, just head over to the Services page and send a request. You never know, I might well be able to save you a lot of time, effort and money.

Amy’s Microtome Counter

Yesterday something awesome happened: Amy (one of the other postdocs in the lab) came to me with an idea for a piece of kit that would help her in the lab. She found that when she was cutting brain sections on the microtome, she would sometimes zone out and forget which plate she was up to. Her request was for a microtome section counter.

The idea was to have something that would light up an indicator of which plate she needed to put the next section into. It would also require a sensor of some kind that would be activated each time she placed a section. We quickly came up with a workable idea that would use an IR sensor, and the user would sweep their hand close to the sensor after each brain section was added.

This project is just asking for an Arduino, so first thing I did was sketch out some code:

// Specify pin connections
int IR = 2;
int LED1 = 3;
int LED2 = 4;
int LED3 = 5;
int LED4 = 6;
int toggle = 7;

// Specify other variables
int count = 0;            // Counter
int maxcount = 4;         // Number of plates to count
long delayTime = 500;     // Delay time to prevent multiple activations
long lastTime = 0;        // Time stamp from last activation

void setup() {

// Specify pin setup
pinMode(IR, INPUT_PULLUP);
pinMode(LED1, OUTPUT);
pinMode(LED2, OUTPUT);
pinMode(LED3, OUTPUT);
pinMode(LED4, OUTPUT);
pinMode(toggle, INPUT_PULLUP);

}

void loop() {

// Toggle to select number of plates being used
if(digitalRead(toggle) == LOW){
  maxcount = 3;
}
else{
  maxcount = 4;
}

// Detection by IR sensor 
if (digitalRead(IR) == LOW){
  if(millis() - delayTime > lastTime){    // Have you exceeded time since last activation?
    count++;                              // Add to counter
    lastTime = millis();                  // Specify timestamp of IR activation
  }
}

// Overflow counter back to start
if (count >= maxcount){              
  count = 0;
}

// Switch on each LED in turn depending on counter
if(count == 0){
  digitalWrite(LED1,HIGH);
  digitalWrite(LED2,LOW);
  digitalWrite(LED3,LOW);
  digitalWrite(LED4,LOW);
}

if(count == 1){
  digitalWrite(LED1,LOW);
  digitalWrite(LED2,HIGH);
  digitalWrite(LED3,LOW);
  digitalWrite(LED4,LOW);
}

if(count == 2){
  digitalWrite(LED1,LOW);
  digitalWrite(LED2,LOW);
  digitalWrite(LED3,HIGH);
  digitalWrite(LED4,LOW);
}

if(count == 3){
  digitalWrite(LED1,LOW);
  digitalWrite(LED2,LOW);
  digitalWrite(LED3,LOW);
  digitalWrite(LED4,HIGH);
}

// Reset timer overflow
if(millis() - lastTime < 0){
  lastTime = millis() - lastTime;
}

}

The electronics is fairly straightforward:

Amy's microtome section counter schematic.

A small amount of assembly later, and we had a working prototype:

Microtome sectioning counter
Amy’s microtome counter

Amy has promised to test it. I’ll let you guys know how it goes.

Stick or Float?

I recently ended my optogenetics experiment (that I showed the planning for in a previous post), and the next step was to cut the brains. As part of the study, I stimulated the mice 90 mins before perfusion, so that I could immuno stain for c-fos as a marker of neuronal activation. This is important for showing that I can successfully activate my ChR2-expressing neurones, but it also allows me to investigate potential downstream sites of activation throughout the brain. All of which is to say that I had 10 brains to cut, and had to cut and save every section throughout their entirety. Which is a lot of brain sections to cut.

A quick mention of my methods, I am cutting the brains to 30 µm sections on a freezing sledge microtome, and placing the sections in phosphate buffer for free-floating immuno. This is not a technique that I had used, or even seen, before coming to my current lab, so my goal for today is to highlight some of the differences between this method and the one I used previously, namely using a cryostat and doing on-slide immuno.

The microtome is a lovely old-school piece of kit, made of massive cast-iron segments, with a cooling stage to freeze your tissue on, and smooth steel rails to run the stage back and forth on the blade. Speaking of the blade, we use modern microtome blades that fit into a holder, but we still have the original chunk of steel that needs sharpening by hand. For those who’ve never used one, it is slightly odd the first time you use it, because the tissue is frozen on the stage, and when you cut a section it melts at the same time and ends up in a little pile of mush on the blade. You then use a paintbrush to sweep it off and into a well of buffer.

Doing the immuno on floating sections takes some getting used to as well, because you need to suction out the buffer for each step of the immuno, but it takes practice to avoid sucking up the sections (water tension is a bitch). But the most challenging aspect of this method is mounting the sections at the end of the protocol, because you now have to half-float half-push your section onto the slide. It takes a lot of practice to do this in an orderly row, without folding or damaging the sections, and without taking endless hours.

By contrast, mounting sections using a cryostat is easy. Because you do all the cutting and mounting in a frozen chamber, each section is frozen and, for want of a better word, stiff. There are two schools of thought for the mounting procedure: I was originally taught do it onto frozen slides, in which case you manoeuvre the section to where you want it on the slide using a paintbrush then using the warmth of your finger on the underside of the slide you melt the section into place. This works well but it does tend to take a toll on the warmth in your fingers (I manage fine because my circulation is great and my hands are always really warm; the PhD student that taught me had to stop every couple of hours after her fingers became too cold to melt the section into place). The alternative is to leave the slice out on the bench, and then you touch them to the section and they melt on in place.

When you mount sections onto the slides as you cut them, you then have to do the immuno on the slide, which makes washes really easy because you can just dunk them into a well of buffer – no suctioning of tissue to worry about. However, it becomes slightly more challenging to do the antibody steps, because you need to do it on the slide, which means placing the slides flat on the bench and pipetting a few hundred microlitres or so of solution to sit on the slide – in this case water tension is your friend. I found that the bubble of solution on the slide is not level all the way across, which means that you end up with a gradient of antibody and therefore the staining is not uniform across your sections. This doesn’t have to matter much, unless what you need is a quantitative staining.

When it comes to planning which method to use for doing immunohistochemistry, I use the following criteria:

  • Use the microtome and free-floating immuno if you want a quantitative readout, such as c-fos, or if you have a lot of brains to cut (I find it’s quicker overall)
  • Use the cryostat and on-slide immuno if you are using a method with very low volumes, such as in situ hybridisation, or you want precise placement of sections

Hey HotSHOT

I was doing a massive round of genotyping yesterday, and was happy to have a rapid DNA extraction method. It reminded of the summer of 2014, when my supervisor had been awarded 3 grants. So we had 3 postdocs due to start at the same time, at the beginning of October. In preparation of the new arrivals, we had acquired a few new transgenic lines and were breeding up a bunch of crosses to produce experimental animals.

This was great news all round, except that come July two PhD students and a postdoc had recently left, and the only other person in the lab was our technician. And we now had hundreds of mice being weaned that needed genotyping. I ended up ear punching a dozen or more cages of mice a day for weeks on end, and with tremendous help from our technician, we were running hundreds of PCR reactions a week.

Ok, so where am I going with all this? The part of this process that took the longest was extracting DNA from the ear notches to use for PCR, and I was getting annoyed by the time-intensive nature of it.

We used a proteinase K digestion overnight, followed by isopropanol precipitation and cleaning with ethanol before reconstituting in water, and it took hours. And we were doing this every day, for weeks. So, I started looking around for quicker and easier methods – I was quite sure we didn’t need such high quality sample prep for a genotyping PCR.

I found a method published in 2000 (and it turns out everyone uses this because it’s so quick and easy, but I had never heard of it), which used alkaline and high heat to degrade the tissue, followed by an acidic buffer mixed in to neutralise the alkali1.

Just in case any readers haven’t come across this method before, I cannot recommend it highly enough. We use it routinely now, and there’s only a couple of PCR’s we run that require a cleaner sample, and then we do our old long extraction.

A quick summary of the rapid DNA extraction protocol:

  • Lysis buffer (for 50 ml)
    • 20 µl of 0.5 M EDTA
    • 125 µl of 10M NaOH
    • 50 ml MilliQ water
  • Neutralising buffer (for 50 ml)
    • 315 mg of Tris HCl (don’t use pre-buffered Tris solution)
    • 50 ml MilliQ water
  • Place tissue sample (eg. ear notch) in a 1.5 ml eppendorff
  • Spin the tissue samples at 13,000 rpm for 30 s to get tissue to bottom of tube
  • Pipette 75 µl of lysis buffer onto sample and place on 95 C heat block for 45 mins
  • Place samples on ice for 5 mins, then briefly spin contents to bottom of tube
  • Pipette 75 µl of neutralising buffer onto sample and vortex

The whole protocol only takes about an hour, and then the samples are ready to use for PCR. Unfortunately, I only came across this method after many many (MANY) hours spent doing our old slow extraction method. It does, however, continue to save us a lot of time in the lab, so I’m happy I found it, and maybe it will help someone else out there save time as well.

1. Truett et al. Biotechniques 29(1), 52-54 (2000) Preparation of PCR-quality mouse genomic DNA with hot sodium hydroxide and tris (HotSHOT).

The CrumbleHopper™

A dietary issue

A colleague approached me a few weeks ago about a laboratory mouse feeder. She was having an issue with one of her experiments; she knew I had done some 3D printing and thought I might be able to provide a solution. Her problem was that she was feeding her mice a special, but very crumbly, diet.

The crumbly diet was making it impossible for her to monitor caloric intake using our currently available food hoppers. Depending on the type of hopper, either the food would crumble all over the cage bottom, or the mice would kick bedding up in with the food and make any accurate food measurement impossible. What she needed was a laboratory mouse feeder that was up to the task.

Designing a mouse feeder

So, my colleague asked me to design and produce a small food hopper that could hold crumbly food without losing it, while still allowing the mice easy access to eat without being able to kick bedding in. It was also important that the hopper be easily added or removed from the cage to allow daily weighing of the food. And as the planned experiment involved some 40-odd animals, they really needed to be low maintenance, and quick and easy to use.

Based on the cages that we have in our animal facility (pretty standard IVC’s from Tecniplast1), I figured the most obvious solution would be to have a hopper that you could hang off the metal grill hopper already present in the cage. Then it would be easily removable as well as being elevated off the floor, and it would be a simple matter to design it to be an easily munchable height from the cage floor.

I “borrowed” a cage from the facility to measure up, and designed what is essentially a hanging basket with hooks. Then after a suggestion from my colleague I made a few variants with different size/shape holes for the mice to access the food:

Initial designs for a laboratory mouse feeder.

Improving the design

I printed these 3 prototypes and gave them to my colleague to test out on the mice. She obviously monitored them very closely, and found that the mice would happily eat from any of them. She did, however, find that the supporting struts were a bit flimsy, so I strengthened those.

Of the three designs, she liked the vertical rectangular slits the best, but wanted the holes slightly wider to allow better access for mouse nom-noms. So, along with a slight adjustment to the height of the hooks to give better attachment, I present the final version of my laboratory mouse feeder, the CrumbleHopper™:

The Crumblehopper: a handy laboratory mouse feeder.

I actually quite enjoyed trying to make these for my colleague, and it does seem to fulfil an otherwise unmet need. So to that end, I am setting myself a couple of goals:

  • To keep an eye open for other experimental situations that could do with having an improved piece of kit. I already have a couple in mind that I am working to produce a solution for.
  • I hope these pieces of lab kit I am developing will be useful to people outside my lab, so to that end I am working towards setting up a small shop on this website to sell the things at a reasonable cost. Where I can, I will also make the designs available online so people with access to 3D printers and such can make them themselves.

Finally, if any of my loyal readers have a niggling problem that could be solved with a relatively straightforward (but otherwise non-existent) solution, please contact me and maybe I can design a solution, as I did for my colleague.

1. https://www.tecniplast.it/uk/product/green-line-ivc-sealsafe-plus-mouse.html