Category: 3R’s

Technology and the 3 R’s

A contemporary approach to the refinement of animal research highlights the importance of technology and the 3 R’s:

“Employing new in vivo technologies that can benefit animal welfare and science including methods to minimise pain and distress as well as to deliver enhancements in animal care, housing, handling, training and use”1

This definition fits well with my experience of animals in research – the less you stress your animals while running experiments, the better your data will be, and technology is an important way to reduce your impact on the animal whilst also improving your metrics.

My history with technology

As an example, for my PhD I wanted to investigate control of the cardiovascular system in a transgenic mouse model. I won’t go in to details here, but suffice to say it had a weird hypermetabolic phenotype which we thought would also impact the sympathetic control of the cardiovascular system. But how best to go about measuring this?

Let’s say you went to your GP with a suspected blood pressure issue, what would the doctor do? Most likely sit you down and measure your blood pressure with a plethysmograph (the cuff that goes on your arm and inflates) and your heart rate by counting beats. This was actually the first method I tried in my mice – we had a mouse-sized plethysmograph that works on the tail of the animal; it also comes with tubes to hold the mice steady while performing the experiment.

You don’t need to be a scientific researcher to realise that confining a mouse in a restricted holder like this will stress them out, even after repeated sessions of acclimatisation. And what happens when you’re stressed? Increased heart rate and blood pressure, which by definition makes this technique less than optimal.

However, it is possible to get decent data from such a system, so long as you bear in the mind the limitations when planning studies and drawing conclusions. As it happens, my transgenic mice that I used in this study were much smaller than the wildtypes, and as such I was unable to get reliable readings.

But, we really wanted to record blood pressure in this mouse model, so next step was to attempt a more invasive method to get a direct reading of blood pressure. This is possible, although technically very challenging in mice, by inserting a thin plastic tube into a major blood vessel and getting a direct readout of the pressure from inside the artery.

Obviously, you can’t go inserting a catheter into the artery of an awake mouse, so I anaesthetised my transgenics and learned how to insert the blood pressure catheter into the mouse’s carotid artery. We did get lovely blood pressure readings from this study, with the predictable caveat that it was performed in anaesthetised animals, and there aren’t many things in this world more likely to impact your cardiovascular system than being anaesthetised!

As it happens, we used a certain (old-school, not used in humans) anaesthetic known to have minimal impacts on the cardiovascular system. However, we knew we wouldn’t be able to publish the results without getting some kind of readings in an awake animal. This is where technology comes into the story, in the form of implantable telemetry.

It goes without saying that telemetry, while a great technological solution, is also technically quite challenging as well as prohibitively expensive. As such, and especially given that I was naïve to these methods, I opted for the easier biopotential transmitter to record ECG. Once I’d figured the experiments out and was able to get good heart rate recordings in awake freely moving mice, they formed the pinnacle of my PhD, and enabled us to publish the results.

In an ideal world, we would have used blood pressure telemetry (heart rate alone can give ambiguous results), but I think we made the correct decision at the time. One of my colleagues recently used the blood pressure telemeters, and had a terrible time of it – they’re just that much more difficult, especially the surgery.

Technology is important

Anyway, my takeaway message today relates the importance of technology and the 3 R’s for minimising the stress and harm done to animals in your experiments while simultaneously maximising the quality and impact of data you produce. I recently submitted a grant application to the NC3R’s with exactly this stated goal – to use my fibre-free optogenetics technology in vivo, to show that it has a marked benefit to animals in optogenetics studies, leading to a refinement, as outlined in the 3R’s.

1. Clark Br J Nutri 120(S1), S1-S7 (2018) The 3Rs in research: a contemporary approach to replacement, reduction and refinement

Feeling Warm and Fuzzy

I have previously talked about developing a touch-free timer for use in surgery. The goal was to better enable a single researcher to maintain sterility during animal surgeries. I really think this is a genuine unmet need in the research world, and widespread adoption of touch-free surgery kit would be extremely beneficial, both to the researchers and to the animals.

Anyway, with the plan to expand my touch-free surgery range, I figured the next piece of kit should be a heat mat for keeping rodents warm in surgery. And again, I wanted something that can be controlled by touch-free sensors. Helpfully, Pi Hut sell a small, flexible heating pad:

Looking at the specs, it uses ~1 A of power, which is far more than we can safely run from an Arduino digital pin. In order to do this, we make use of a component called a MOSFET, which is a special kind of transistor used to amplify circuits. A MOSFET lets you use a digital signal (eg. an Arduino output pin) to fully switch a separate circuit (eg. a fully powered heat mat).

Therefore, using a MOSFET, I can control the power going to the heat mat by the digital output of the Arduino. I’ve mentioned pulse width modulation (PWM) before, and it is perfectly suited to this application. PWM will let me digitally control the amount of power going through the heat mat. And, best of all, because it’s digitally controlled, I can shift the PWM up/down with IR proximity sensors.

But, how to display the power going through the heat mat? For this, I again turned to Pi Hut, who sell a 10-segment LED bar:

Each LED in the bar is individually controlled, which means that I can set the Arduino to display an indication of the power going through the heat mat, on a scale of 1-10. Bringing it all together in a 3D printed housing, I have power up and power down proximity sensors, a power indicator bar, and a flexible heat mat that warms quickly to the extent determined by the user:

Touch-free heat mat for keeping rodents warm in surgery.

I have used this heat mat in surgery myself, and it worked really well. It heated up super quick and I could change the power of the heat mat to the temperature needed by the mouse. This piece of kit is indispensible for keeping rodents warm in surgery.

The one thing that I think it missing is an actual reading of the mouse’s temperature – I kept having to feel the surgery bed to check the temperature, which kind of defeats the purpose of being touch-free.

So, my next plan for this piece of kit is to add in a temperature sensor (whether a standalone one or one that runs through the Arduino, I have yet to figure out). Stay tuned for updates.

A Better Timer

This is a quick update to my previous blog post, where I explained making a touch-free surgery timer. Unfortunately, I found that it just wasn’t working well – as I explained previously, the low voltage used by that timer (1.5 V from a single AA battery) was not enough to make multiple IR sensors respond to the proximity of a finger.

After some brain-storming for how to fix this issue with sensing, I came to the conclusion that I really needed to start from scratch with a new (higher voltage) timer. I therefore scoured the internet for a timer with the following features:

  • Runs at 3-5 volts (ie. two or three 1.5 V batteries), but not from button batteries (they don’t have much capacity, so would drain too quickly)
  • Stopwatch and countdown timer functions
  • Controlled by limited number of pushbuttons (ideally Start/Stop and Reset for stopwatch, then “add minute” button to set up timer)
  • Has big light-up numbers for easy checking of timing

Finally, I found a timer that hit all my requirements, and only cost around £20 on Amazon (turns out the kind of timer I wanted with simple controls and big light-up numbers is targeted to children and the elderly):

So, I cracked the timer open, soldered in IR proximity sensors to the Start/Stop, Reset and Minute buttons, and found they worked great – I could control the timer by moving my finger to within 10 mm or so. Turns out there was some dead space inside the case, which meant that I could drill holes in the side and mount the sensors on the inside. I also added in a cutoff toggle switch to cut the power to the IR LED’s to prevent battery drain.

All in all, I’m really pleased with the new timer: I left it in the surgery room, and it’s going down brilliantly with the other lab members who’ve tried it.

How to Track a Mouse

Our old locomotor tracking

One of my projects is investigating a population of neurones that controls mouse locomotor activity and food intake. In the past I have used either implantable telemetry or IR beam break cages to quantify the mice’s movement. But the telemeters, even when they’re functioning well, don’t give particularly good quantification of mouse locomotor activity, which leaves the beam break cages.

For anyone that doesn’t know, these cages are set up to have a couple of IR beams that cross the cage. Whenever the beam is broken (ie. the mouse gets in the way), this is registered by the computer. It’s quite an effective (although crude) method to quantify mouse activity. And it does so completely non-invasively. However, our current IR beam break cages have a number of drawbacks that make them unattractive:

  • They only work with some of the older open cages, and don’t work at all if the mice have any bedding in the cage (it blocks the beams)
  • The beam break cages we have available in the facility, which actually belong to one of the other lecturers (although she is happy for us to use them), are a decade or two old and were built by a previous postdoc – as such they have suffered some degradation over the years and only have partial functionality left

Anyone who reads my blog will already know what I’m about to say – with these issues I’ve raised, I decided to try and build my own set of beam break cages.

Setting up beam breaks

Right, so first step was to find some IR LED’s and sensors that I could pair across 20-30 cm of a mouse’s cage. I’ve used things like this in the past, so I know you can detect an IR signal using an LED in the ~900 nm range and a phototransistor (Figure 1A).

Luckily, I had some sat around, so I hooked them up to an Arduino, but could only detect the IR signal up to around 5 cm distance. This is obviously not enough, so after some detective work, I found some “IR Beam Break Sensors” from PiHut (Figure 1B). If those didn’t work, it would require some more complex electrical engineering to make it work. Apparently you need to use modulated signals to be sensitive enough to work over multiple metres.

Fortunately, the IR sensors from PiHut worked a treat, up to about 40 cm, which is more than enough for my purposes. The next issue was how to fix the sensors in a way that they would remain aligned in a pairing across the cage.

Aligning the sensors

For this I turned to my trusted 3D printer. After borrowing an IVC from the animal facility, I figured I could make hanging holders that would hook onto the side ridges (Figure 2).

These worked great, with the only issue that the mice tended to move their bedding around and block the direct beams. A very simple solution to this problem was to use strong neodymium magnets to “pin” the tube/bedding at one end of the cage, out of the way of the sensor beams.

Right, so now I had 2 pairs of sensors successfully attached to each mouse cage, next I needed to actually track the data in some way.

Tracking data using Arduino

It turns out that tallying IR beam crosses is easy peasy using an Arduino. The only annoyance being having to duplicate the code 24 times (ie. 2 sensors each for 12 cages). But, I still need to get the data out of the Arduino. I figured I could either hook up an SD card reader and write the data to a removable card, or hook up to a PC and download the data directly.

As I was already connecting the Arduino to my laptop, I tried that first. A little Google sleuthing found me an open source (ie. free) “terminal” program, that will happily log data that comes in over a “COM” port, such as is used by the Arduino. It was actually really easy to set up, and will log the IR beam break data in a CSV (comma separated values) format, that can be directly opened by Excel.

For ease of later data analysis, I made the program log the data in 10 second intervals. However, it will be easy to change that depending on the experimental paradigm eg. 1 or even 10 minute intervals for longer term studies over days or weeks.

Just to prove how well the system works, you can see a massive increase in activity following injection of caffeine (Figure 3A). You also get fantastic circadian activity if you record for longer time periods (Figure 3B).

Where to get it from

As always, I am making this system available on my shop, far cheaper than any commercially available system. Obviously I’ll include a copy of the data logging software with instructions of how to use it. Anyone who wants to measure mouse locomotor activity easily and cheaply, check it out.

Edit 5/5/22: I have now uploaded details of how to make this kit to Hackaday, so head over there if you want to try and build it yourself.

Doing Away With Fibre

My interest in wireless optogenetics has come up a couple of times. In fact, I’ll start with a quick correction: I prefer to call it fibre-free optogenetics, after multiple people mistook my wireless system I was designing as meaning controlled via Bluetooth or WiFi. Which it ain’t. And, for me at least, the whole point of going “wireless” is to do away with the optic fibres, which really embody all the issues and difficulties with in vivo optogenetics:

  • Impacts to the animal – the need to have the animals in an open cage, with an open lid and a sterile environment to prevent damage to the fibres. Also, they tend to be stiff, having severe behavioural impacts.
  • Loss of optical power – the optic fibres require additional optical connections, which inevitably leads to light loss, and therefore difficulties obtaining a high enough brightness.
  • Expensive and fragile – not much more to say, other than we have spent thousands of pounds maintaining the optic fibres for our optogenetics system. This may be more than is typical, but I think that’s because the Plexon fibres we use are very fine and lightweight – I have used more durable ones that were even worse for the mouse behaviour because of the added stiffness.

The most important reason to do away with the optic fibres, as far as I’m concerned, is the impact to the animal. Quite apart from minding the 3R’s with regards to animal welfare, tethering will inevitably cause stress, which is detrimental to the data you can acquire (Figure 1). In fact, it is to the NC3R’s that I am applying for funding to take my fibre-free opto system to the next level.

There is of course the added bonus with wireless optogenetics that you can do optogenetic stimulation in otherwise impossible setups. For example, I am very keen to use my fibre-free opto’s in our calorimetry system to measure energy expenditure in response to opto stim. This is done in an air-tight sealed container, which to my knowledge this has never been done with optogenetic stimulation in the brain.

After a fair bit of research, I have found 4 commercially available wireless in vivo optogenetics systems (Figure 2).

Helios by Plexon and Teleopto by Amuza are both very similar, except that the Helios headstage attaches to “normal” implants, whereas Teleopto make their own custom implants. Both require you to point an IR remote at the headstage constantly (ie. the flashing stops if the signal stops). Fi-Wi from Doric connects over radio signal to drive opto flashing; similar to Teleopto they use custom implants. Neurolux is a very different system to the other three, and uses electromagnetic induction to remotely power the implants. Hence the Neurolux implants are tiny and custom (the LED is actually on the end of the fibre that gets implanted).

I have collated a summary table of the various systems, including a number of parameters (Table 1). Included is the cost to buy a complete setup to stimulate 1 mouse at a time, which usually comes with a few implants. However, I was unable to find out the irradiance available from the Plexon Helios system, despite asking the sales people for those details.

Overall, the Doric system seems the best of the bunch; despite being the heaviest it is very compact and produces by far the highest irradiance. In fact, it provides higher irradiance than the system I’ve been developing, which comes out around 150 mW/mm2. Stay tuned, and I’ll be talking more about my system in the coming months.

1. Won et al., Nat Biomed Eng (2021) Wireless and battery-free technologies for nanoengineering.

No Touchy!

This came about when I was thinking about how to improve my surgery technique, in particular sterility. Not that I or people I’ve taught have ever had issues with infection, but maintaining good sterile technique can be difficult, and it certainly makes many routine tasks challenging. And, unlike “real” surgeons, who have a team of underlings to aid in their surgery, we researchers often undertake ours alone, or at most with a single technical helper.

So, what kind of routine tasks do we perform during surgery that could be done without touching anything? My first thought was a timer – we do a lot of virus injections with a nanoinjector, and we time how long we inject to ensure sufficient spread of the virus before retracting the needle. After much Googling and scouring of the internet, I was unable to find any “hands-free timer” that wasn’t a 30-second hand-washing timer. So I decided to develop my own.

First step to a hands-free timer is to find a switch of some kind that can be remotely activated, a proximity sensor of some kind. I wanted something cheap and crudely effective – ie. something that triggers upon proximity of an object (likely a human finger), with a detection distance of 5-10 mm. This lead me to two likely alternatives, a capacitive plate or an infrared sensor (Figure 1).

I bought some sensors to test, hooked them up to an Arduino and found that the IR sensor worked great, but the capacitive plate was very hit-or-miss. Right, so I knew which sensor I wanted to use, but now I needed to set them up on a timer. My first thought was to use my perennial favourite, the Arduino. And I did start setting that up, but then I found an old lab timer that I thought might make a quick and easy prototype (Figure 2A).

Opening the old timer up, I found that the buttons were simple conductive latches that connected the microcontroller pin to ground. All I had to do was correctly solder in the phototransistors of my IR sensors to perform the same latch, and I could activate the “buttons”. The one snag I found was that the timer ran off a single 1.5 V battery, which wasn’t really enough to produce a strong IR signal. This meant a couple of things:

  • I could only connect two IR sensors before the drain on the battery meant they ceased to function entirely; therefore, I hooked them up to START/STOP and RESET, to give me a functional stopwatch without the other functions.
  • The low power available from a single AA battery meant that even with only 2 IR sensors connected, they have low IR strength and therefore can only detect highly IR-reflective objects. It works for metal objects and gloved hands, so should work fine in surgery.

I connected up a couple of IR sensors to the existing circuit, as well as a toggle switch to turn off the power to the sensors to prevent battery drain when not in use. I then 3D-printed a box to house everything, giving me a prototype for my Touch-Free Timer (Figure 2B).

I had assumed that the timer, being so old, would not be in production any more. However, it looks like you can still buy it from lab suppliers, so I shouldn’t have any problems producing as many as are needed.

Animal Consumption

A shower thought

I was having an imaginary argument this morning – you know, the kind you have in the shower where all your points are zingers and your opponent can only be floored by your insightful oratory, whereas anything they come out with is antiquated and flawed. On this occasion, my imaginary antagonist was my father-in-law, who is great for such things because he is a classic dogmatic conservative who apparently changes his mind only when instructed to do so by the Daily Mail. He is also loud and steamrolls all other voices in his vicinity, such that my wife is the only person who successfully argues against him.

Anyway, on this occasion, I was actually walking to the train station, which is another great time for introspection, when I started thinking about the recent news that South Korea’s president was considering banning the consumption of dog meat. Now, I could just imaging the FIL lauding this in his typical brash manner: finally some sense, how could this culture engage in such a disgusting practice for so long?

Animal lover

Now, for context, my FIL absolutely loves dogs, so this is a) a very reasonable position for me to give his fictional self, and b) not something I would ever argue with him in real life. But, as this is a fictional confrontation, there’s no problem. So my rebuttal would go along the lines that, yes I agree that eating dogs is distasteful, and not something that I would ever even consider doing, but how is it any different from our consumption of pigs, cows and sheep?

There is no argument you can make against the consumption of dog meat that doesn’t also preclude the eating of any animal without resorting to playing to our cultural history of keeping dogs as pets. And at that point, one can just point to the historical culture of eating dog meat in places such as Korea.

Pigs in particular are as sociable as dogs, and at least as clever. I’ve seen videos of cows bounding around like puppies and showing affection to their owners, and I know people who keep chickens (and other birds) as pets. The same could be true of rodents, not that we eat them, but we do exterminate them fairly indiscriminately, and I can testify that rats are both clever and sociable. Horses and dogs are often used as working animals (not that I enjoy eating horse meat, but there is a historical precedent of them ending up in food as a cheap substitute to beef).

Advocating veganism

This argument inexorably leads to advocating veganism, which my wife and I attempted a couple of years ago, but found it too challenging; if you ever check the ingredients of packaged food, 99% of the time it contains animal products (particularly dairy), even in food that you would never think it necessary. Instead, we went for drastically reducing our consumption of animal products and when we do, making ethical choices.

We have swapped out cow’s milk for oat milk (which is a bit more expensive but I actually prefer it), only buy free-range eggs (which we did anyway), and try to buy ethically produced meat on the occasions we do buy it (probably once a week). Unfortunately, I am a total cheese hound, which has been the hardest thing to cut.

Extending the argument at the other extreme, how can you argue against consuming any animal? I remember watching a program about people (poachers, I guess?) hunting and eating wild animals in the jungles of Borneo, which went by the deceptively innocuous term bush meat. As this is Borneo, every animal is likely to be in danger of going extinct, which makes it easy to vilify and argue for a total ban. But, as a middle class person who’s grown up in wealthy nations and never been hungry or homeless, how can I judge people for hunting for food?

As I said earlier, many of the ethical changes I’ve made to my diet also increased the cost, so how can I judge others who don’t have the means to do so? Well, when they show that sometimes “bush meat” includes orang-utans or chimpanzees, suddenly my sympathies evaporate. And finally, we come back to the arguments in the use of animals in research – balancing need against ethical usage and suffering.

Reducing suffering

A basic criterion is the ability of the animal to feel suffering, which increases with the innate intelligence of the animal, which is why we are so instantly disgusted by the suffering of primates. And also why a huge amount of animal research is performed on mice, who sit in a good balance between less capable of suffering but close enough to humans to enable important and relevant research.

At the end of the day, reducing the suffering of animals around the world comes down to two things: education (about the harm being done; eg. see the NC3R’s) and empowerment (particularly financial, to enable change). This is particularly true when it comes to eating animals, where we can obviate the need for animal consumption, but only with a huge and concerted effort.

The 3 R’s

Replacement. Reduction. Refinement. Also known as the 3 R’s.

On the face of it, the 3 R’s form a fairly straightforward guide to limit the amount of suffering endured by animals in your experiments. However, these are stepping stones to quite indepth process for advancing technologies and rigorous planning, as defined by the following on the NC3R’s website1:

  • Replacement – Accelerating the development and use of models and tools, based on the latest science and technologies, to address important scientific questions without the use of animals
  • Reduction – Appropriately designed and analysed animal experiments that are robust and reproducible, and truly add to the knowledge base
  • Refinement – Advancing animal welfare by exploiting the latest in vivo technologies and by improving understanding of the impact of welfare on scientific outcomes

These are major points that I think about frequently when planning and performing animal experiments.

Before you plan an experiment

Replacement would seem quite straightforward for someone who works on the mouse neural system, in that it’s not something in my control, so not to worry about it. And while it is true that I rarely have intentions to work in non-animals systems, that doesn’t mean it’s irrelevant.

Really, this needs addressing at the most fundamental level, before I even plan an experiment, ie. is the scientific question I want to answer relevant to a whole-animal neural system. Does it require the use of an animal to answer this question?

For example, I have often used relatively unknown neuropeptide agonists in my work; if I wanted to know more about the intracellular signalling mechanisms these agonists use, it would be both unethical and a waste of time, money and animals to test this on live brain slices (which I use for patch clamping). Instead, one would use a cultured cell line, such as HeLa cells.

Robust and reproducible

Reduction is an interesting one. It’s easy to think, well I’ll just use fewer animals in my experiment. However, this misses the key point of “robust and reproducible” experiments. What if you used fewer animals and didn’t see an effect? Is that because there is no biological effect of your treatment, or is it because you didn’t have enough animals in your study to show a statistical effect? This is where power analyses come in to play: they help you plan a robust study without using an unnecessary number of animals.

It is also important to think of optimising your study design to produce the most statistical power (eg. using crossover studies and repeated measures ANOVA) and to negate the need of repeating studies in the future. Even during an experiment, I am conscious of this metric, because I am always trying to reduce the variability in a study (for example, by reducing animal stress) in order to improve the power and numbers needed for future studies.

Improve data quality and impact

Refinement is really where it’s at for me. I spend a lot of time optimising techniques and, more recently, developing technology, to improve experimental conditions. The thing is, when your mice are unhappy and stressed, they will not behave naturally and your data will be more variable. So it makes sense, from both a pragmatic point of view and for animal welfare, to refine your experiments as best you can.

Refinement can include anything from your study design, acclimatisation of the animals and their housing conditions to advances in technology allowing better data to be collected.

Technology and the 3 R’s

Ever since my PhD, I have been interested in the use of technology to produce better data, and improve animal welfare along the way. I was particularly keen on the use of telemetry to obtain high quality physiology data while minimising the stressful environment. Since then, I’ve been interested in using AAV’s to target neurone populations of interest, and then more advanced technologies including optogenetics and fibre photometry.

In addition to improving the animal welfare in a single experiment, these more advanced technologies can provide more impactful data with deeper insights, which means fewer studies need to be performed to provide a clear picture of the biology in question.

I have also become very interested in developing in vivo technologies myself to improve on aspects that I know impact negatively on animal welfare, for example trying to perform fibre-free optogenetics to limit a lot of the negative aspects of those experiments (such as the need to have head-tethered animals in open cages during experimentation).

A 3 R’s framework

The 3 R’s provide an excellent framework with which to approach animal research in a way that aims to be as ethical as possible. And in fact, I would argue that we are morally bound to consider such questions whenever we intend to perform experiments on animals.

1. https://www.nc3rs.org.uk/the-3rs

Animals in Research

“What do you think about the use of animals in research?”

This is the one question you can always guarantee will be asked in a job interview that has anything to do with the use of animals in research. And it’s actually quite a difficult question to answer well. The answer clearly lies somewhere between, “Yeah, I’m fine with it, I don’t care” and “We have no right to use animals like that for our own benefit.” But, how do you justify the experimentation of animals without coming across as glib or self-serving?

Quite apart from job interviews, I’ve always found this to be a difficult topic. I guess because it is both emotive and there is so much misinformation surrounding it. Generally, the only time you hear about animal research in the news is when there’s been a particularly horrific protest. And those who are most able to talk about the reality of animals in research, ie. the researchers, are scared and drilled into not speaking about it. To the point that only my family and a couple of close friends know what I actually do for a living.

Protesters against animal research.
Anti-vivisection protest in the US

Almost 75% of animal research is performed on mice

I will occasionally, once every year or two, come across an anti-vivisection “information” stand at a market or something, where they are handing out leaflets to try and persuade the public as to the evils of animal research, and the pleasure the scientists take from doing horrific things to the animals. This is obviously not the case, and it’s somewhat indicative of the weakness of their position that they have to cherrypick and inflate the stats, focussing on the most photogenic species, like dogs, cats and primates.

They completely misrepresent the fact that the vast majority of animal research is performed on mice (Figure 1), and more than 90% in mice, rats or fish.

In fact, there are only a couple of facilities in the country that perform research on monkeys, and in fact all research with chimpanzees and other great apes was stopped in 1998. Speaking of which, the reason all the pictures they use always look so terrible, is that they are all ancient (mostly from the 80’s or earlier, before the regulations were brought in).

Experimenting on animals is difficult and expensive

I’m not sure how many of the public actually know that you need a license to perform scientific research on animals. I say a license, when in fact you need multiple: a personal license for the researcher actually performing the experiments, a project license for the person (usually professor) in charge of the work, and a site license for the location the experiments will actually take place. And everything needs to be justified and planned beforehand, with expected outcomes and experimental group sizes. Then you need named training and competency officers, a vet, animal welfare officer and the technicians who will actually be caring for the animals.

And we mustn’t forget the home office inspectors who can, and will, drop by to check on the welfare of the animals, and make sure all paperwork and training is up to date. All of which means that experimenting on animals is difficult and expensive, and requires huge amounts of training and expertise. So anyone who thinks that overworked and underfunded scientists, quite apart from the moral and legal implications, will be frivolous with their use of animals, is deluded.

How to combat misinformation

There are, of course, institutions trying to combat the spread of misinformation, such as the National Centre for 3 R’s research (www.nc3rs.co.uk). It is, however, very difficult to get the public interested in science and statistics when compared to emotive pictures and moral outrage. Which is why it is so important for those that know better to spread good information about this topic.

But where do you begin, when the scientists have been conditioned to be silent about anything to do with vivisection, and the public are so conditioned to fear the evil scientist? It really has to come down to education, both about the realities of animals research – they are treated far better than farm animals, but the moral outrage clearly lands heavier on experimentation – as well as the benefits to medicine and society that come out of this research.

I’m not going to lecture my readers about all the great advances coming out of animal research; suffice it to say that any medical advance you have ever heard of was borne on the back of a huge amount of scientific research, much of it requiring the use of animals. And by the way, this includes many benefits to modern medicine that people may not think of. So even if there are people out there who take a moral stand and refuse any kind of medication because it required vivisection, you live in a world without smallpox and polio (and, might I add, with a Covid19 vaccine) thanks to the use of animals in research.

One of my goals with this website and blog that I have started is to help spread actual and interesting information about research involving the use of animals. Now it is up to us to be thoughtful and diligent with our use of animals, and make sure their sacrifice is not wasted.