Optogenetics - Science method

I think Matthew and Elias are both answering the question, but from different perspectives.

Matthew is talking about activating neurones eg. with ChR2; then you’re looking at (relatively) fast pulsing on the order of 10 Hz. The goal here is to drive action potentials at the rate/pattern that they would fire naturally. You need to be careful not too pulse too quickly, as that also leads to inhibition in the same way Matthew explains for prolonged activation.

Elias is talking about limiting the light damage, which is particularly relevant when using inhibitory opsins. In this case, the ideal situation would likely be continuous light stimulation, but like Elias says that will lead to tissue warming and cellular damage. In theory, you could drop the light power to limit damage, but then you will drop below the activation threshold for the opsin and nothing will happen. Therefore, we do slow pulses (on the order of 0.1-1 Hz); for example, light on for 5 seconds, then off for 5 seconds. This compromise provides prolonged opsin activation while limiting tissue damage. Be careful not to do fast pulsing of inhibitory opsins, as you can induce reflex action potentials.

I have written about this exact issue on my blog, feel free to check it out and hopefully it will help answer the question in more detail:

It's very odd that in your failed experiments you are getting *literally zero* expression of the AAV as opposed to weak expression (scattered red cells). Do you have records about which mouse was injected on which day? You say you did 2 mice per day and you have 6 mice with expression and 4 with no expression - do those 4 failures represent 2 days on which zero animals worked? Because if that's the case, I think it makes sense to consider things like the virus having been inactivated/destroyed, or a major problem with the equipment setup.

The other thing that comes to mind is the fixation and imaging/detection. Did you perfuse these mice and then post-fix in formalin, or go straight to drop fixation withour prior perfusion? Also, are you imaging the mCherry that is present in the tissue, or using antibody-based detection to add another fluorophore onto it? Consider possibilities like major difference in fixation quality/time (= more protein denaturation = loss of endogenous fluorescence) or photobleaching (could any of these slides have been left out exposed to light for overnight or longer?) Finally, is there any possibility that there was an issue with the imaging step? (Did you swap from a slide that had mCherry on it to a slide that should have had and didn't, to directly confirm that the microscope settings that had just worked to visualise mCherry saw nothing?) I ask because I have done SO MANY things like leaving a shutter closed or the wrong filter set in place...

Many studies have already shown that stimulating axons expressing rhodopsin is susficient to altered the circuit-mediated behavior. I can't recall any study doing dendritic activation. My guess is that if dendritic stimulation is strong enough to activate somatic firing, you should observe behavioral consequence if these neurons are critical.

I can say expression of the Gcamp6s is higher with Ai162 mice vs AAVs.

Hi Simon,

I have not experienced this myself, but I can offer some ideas for troubleshooting:

Some more context would help. Is this the first opto experiment you've done? Is the optogenetic equipment new? Has it been used successfully before, or even validated?

If you've not already done it, I would suggest finding a good paper where someone has done what you are seeking to do, and copy their methodology exactly - same equipment, same cells, same construct in the same serotype virus etc. Also check you are getting decent light power on your sample.

Hope this helps. Feel free to message back if it's still not working.

Nic

Hi Shashank,

I hope I'm not too late to offer my help to your question. I have been putting together an Optogenetics Guide to try and answer exactly your question (and others relating to in vivo optogenetics):

Please do check it out, and hopefully it will give you a good idea of where to start.

Nic

Hi Sertan,

In mice the 1.25 mm ferrule is great. The 2.5 mm ones are really big, and you don't gain anything for using them in mice. Even with the 1.25 mm ferrules, it is difficult to do bilateral stimulation without angling the fibres - you need much more than 1.25 mm separation. If you're not sure how to plan angled fibres, I have developed an online tool to calculate angled stereotaxic coordinates: https://nicneuro.net/calculate-cannula-angle/

As for core size, I would also recommend sticking to 200 um. As others have stated, increasing the diameter will increase the absolute amount of light coming through the fibre. However, it will also disperse the light across a much wider area, so it won't necessarily help the irradiance, which is the critical value for activating opsins.

Finally, always use 0.22 NA fibres (even with LED's), as the higher NA fibres scatter the light too wide, with only a small increase in light output. Overall, your effective stimulation distance is reduced, check out my Optogenetics Depth Calculator for more info: https://nicneuro.net/optogenetics-depth-calculator/

Actually, one more thing: I would advise getting stainless steel ferrules. I found the ceramic ones seize up with the ceramic sleeves, and I ended up ripping some fibres out accidentally. Nowadays I just get these fibres from Thorlabs in boxes of 20 and cut them down to size myself: https://www.thorlabs.com/thorproduct.cfm?partnumber=CFML22U-20

Hi Yannik,

There's a lot to unpack here. Just to clarify, are you talking about the fibre optic patch cables (connecting the LED to the mouse) or are you talking about the optic cannula (that's going in the mouse's brain)? From your screenshot, I'm assuming you mean the patch cords. Doric do have some further info summarising the different types of fibre cores: https://neuro.doriclenses.com/collections/fiber-optic-cannulas/products/mono-fiber-optic-cannulas?productoption%5BFiber-optic%20Material%5D=Silica%20%2F%20Silica&productoption%5BReceptacle%5D=ZF1.25&productoption%5BFiber%20Tip%5D=FLT

Firstly, what optic cannula are you using? For optogenetics, I always recommend 200 um core 0.22 NA fibres, even when using LED sources. Reason being, as soon as you increase the NA of you fibre, you can get a marginal increase in light entering the mouse's brain, but it scatters far more widely, so it's actually less effective. For further info, check out my Optogenetics Power Calculator: https://nicneuro.net/power-calculator/

Next, about the patch cords. As Norman said, the optimal patch cord will depend on your LED. However, assuming that you are using a 200 um optic cannula, you do not need the full 1 mm diameter core. So, my suggestion would to drop the diameter of your patch fibre (maybe to 400 um), to maximise irradiance capture from the LED. In this case, the NA of the fibre shouldn't matter too much, so long as you do use the 0.22 NA fibre cannula.

Lastly, does your patch cable have a rotary joint? If the mouse is head-fixed that isn't necessary, and will lead to an avoidable loss of light power.

Unfortunately, many of the LED's sold for in vivo optogenetics are simply not bright enough. In particular, I'm not sure I've seen any outside of the blue 450-470 nm range that are really bright enough for normal optogenetic tools.

Thank you, Nicolas, I think Pulsepal will help my experiments.

I had something similar when trying to make fibre optic cannulae - the epoxy was just not setting properly. Either that or they were to viscous to get the epoxy into the cannula.

My solution was to stop trying to make them myself. Now I just buy them in pack of 20 from Thorlabs, and they cost less than £20 each. These are the ones I get:

Is there a particular reason you need to make them yourself?

Could you not just transfect your cells with two viruses? Also, I'm not a fan of the hM4Di, it just never seems to really do much, no matter how good the transfection (and it's possible this is caused by accumulation in the cytoplasm https://blog.addgene.org/tagging-optogenetics-and-chemogenetics-receptors-fluorescent-proteins-and-other-options).

There is an alternative opsin that can produce activation or inhibition of a neurone depending on the wavelength of light used. I've never used it, but it looks good: https://www.addgene.org/154951/

I'm going to buck the trend here and suggest you do not use such high NA fibres, regardless of your light source. The problem is that any increase in NA makes the light coming out the end scatter more, decreasing your effective stimulation distance. With a higher NA fibre you need ridiculously high light output to maintain your effective stimulation depth.

For example, in a typical experiment with a 200um fibre and requiring an irradiance of 1mW/mm^2 to activate ChR2, if you wanted to have effective stimulation for 1 mm from the end of your fibre, you would need approximately 4.6 mW from a 0.22 NA fibre. But if you used a 0.65 NA fibre, you would need 27.4 mW. I ran these calculations based on the Aravanis model, and using my online optogenetics power calculator at: https://nicneuro.net/power-calculator/

My experience, using both Plexon and Prizmatix LED's, is that increasing the NA of your fibre only gives a small increase in light power. For example, I recently tested 0.22 NA and 0.50 NA fibres with a Plexbright blue LED and achieved 7.4 and 11.0 mW, respectively. In this case, increasing the NA produced a 50 % increase in light output, but due to the massive increase in light divergence gave an approximate 30 % DECREASE in effective stimulation distance in the mouse brain.

Unfortunately, any system you use to model light propagation in the brain will be an estimate based on experimentally derived scattering values. And the scattering values are wildly different depending on who's done them (anything from around 9 to 40 mm-1).

A Monte Carlo model seems to be the best, but there aren't any simulations I can find that are easy to get quantifiable numbers from (as you seem to have also discovered - I have also used OptogenSim and it provides at best a very rough indication of light spread).

I have developed an online tool (https://nicneuro.net/optogenetics-depth-calculator/) which uses the Aravanis model. You select low, medium or high scattering tissues (depending on the relative amount of grey or white matter), input the parameters of your optogenetics system, and the calculator will give you an estimate of the depth you can expect to activate your opsin.

The important thing to keep in mind is that any light modelling tool will be an estimate, so I would advise using them as a guide to help decide your fibre placement relative to your desired site of activation.

A possible in vivo solution would be to activate your neurone terminals (I like to do a solid 20-min stimulation), and then perfuse your mice 90 mins later and do immunohistochemistry for c-fos in the ChR2-expressing cell bodies. If you compare stimulated vs unstimulated, you should get at least an idea of whether you have activation of those cell bodies. It's not perfect, but it's quite easy to do and well-accepted in the literature.

I usually count all cells in the expressing region and calculate the average cell density over the expressing region. Then you can use some calculator like http://web.stanford.edu/group/dlab/cgi-bin/graph/chart.php

to calculate effective power density in the tissue. If you know the electrical properties of your cells of interest you can approximate the power necessary for efficient modulation.

I can send you a chapter we wrote about how we do it. DOI:10.1007/978-1-4939-7417-7_8

Dear Simon Miguel Lopez ,

That's the least I could do.

Stephen David Edwards & Bhogaraju Anand,

Thank you for your answers.

As suggested above, you should try lower volumes and injection speeds. Also, you could aim to the lateral portion of the BLA if leakage into the CeA is your main concern.

Another thing to consider is how you define a "leak"? If you are simply looking at fluorescence intensity with a relatively small magnification then you might mistake labeled axons coming from the BLA as a leak. This is not so trivial to solve with some vectors, if the fluorophore is directly attached to the opsin (and therefore does not fill cells, but rather outlines their membrane). Nevertheless, as a first step, I would suggest staining for NeuN and then using either a confocal or high magnification in a fluorescence microscope (e.g., 20x) to see if it is in fact the CeA somata that are expressing your opsin, or simply a high concentration of opsin-expressing axons from the BLA.

Dear Joe, as an inorganic chemist I'm clearly not an expert enough to give you a qualifed answer to your interesting technical question. All I can do right now is suggest to you the following potentially useful article which might help you in your analysis:

This paper is freely available as public full text on RG. Please check particularly the "Materials and Methods" section on page 2.

Good luck with your research!

I have an update for you guys.

We are using ChRR - Syn now. I thought it could be something about the promoter (CAG) but it doesnt seem so. It's happening again

The pic is a neuron transfected with lipofectamine 2000 and ChrimsonR-Syn at 2:1 ratio and it's clearly dying. However it seems that the cells last a little longer in a "healthy" state with ChRR-Syn.

Best Regards!

Hi Andrea,

We haven't actually tested different intensities for the optogenetics stimulation. My understanding of photobleaching however is that it is a continuous decay in the signal. In these experiments we deliver optogenetic stimulation for 24 seconds and then it turns off. The decrease occurs during those 24 seconds, then the signal rises back up.

I've experienced the same thing, no activity all. But the neurons appear healthy and in perfect morphological state. You are not alone... I've also co infected with Gcam and Chrimson, and the same thing....

I hope I'm not too late to help you with your issue. We also use the Plexon optogenetic system, and have encountered the same issues as you. The good thing about the Plexon system is the fewer optic connections which results in less light loss and a strong light power out the end of the cable. As you say, the commutators are problematic and their optic cables are very fragile, though overall we've found it to be ok. Here are some things we do:

Unfortunately, there are limited options for in vivo optogenetics without the use of lasers (which we wanted to avoid for safety reasons). Out of the LED systems, I think Plexon does make a decent compromise between usability and light power outout, whereas I'm not sure what the actual light output of the other LED systems are, as the manufacturors are reticent to publish that information

@Justin Siemian ,

Thank you for your advice! I will check that out as well.

Hi Soledad,

depends on the optogenetic actuator. If you use an excitatory one this will lead to the release of the neurotransmitter of the affected cell. However if you use an inhibitory one (e.g. Halorhodopsin or Archaerhodopsin ) you hyperpolarize the cell and therefore supress its firing and transmitter release.

I hope this is of some help.

I would look around the new addgenen database

http://datahub.addgene.org/aav/

Hi there!

Recently I solved the trouble!

As you mentioned before, the filament structure was dendrite!

I changed the stimulus source as 4-ap (potassium blocker) induced seizure model.

And finally found that the previous optogenetic stimulus was not enough to induce action potential!

In 4ap model, ipsilateral site had less c-fos positive cell body (may be spontaneous firing) shape but also shows the dendrite structure.

On the other hands, the contralateral site showed a lot of c-fos positive cell body shape near the 4-ap injection site and dendrite structure.

In my opinion, the c-fos antibody which I used was low affinity or weak secondary binding thus I observed the dendrite.

However when the certain stimulation was delivered, the c-fos antibody was enough to distinguish between stimulated and non stimulated neuron.

Now I would change optogenetics stimulation parameter!

Thanks a lot !

you will need to ensure that the excitation spectra of both the indicator (voltage or calcium) and the opsin are sufficiently separate so that you don't have any crosstalk, i.e. the imaging wavelength exciting the opsin.

Our lab found a solution using Chronos for the opsin and CaSiR-1 for the calcium indicator

Most definitely. For example, we and others express ChR2 (or derivative like chronic) in prefrontal cortex and after letting the neurons express for 4-6 weeks, make slices and can optically evoke TTX sensitive monosynaptic EPSCs in brainstem targets (e.g. dorsal raphe). These EPSCs can also be reinstated following TTX blockage by adding 4-AP which, by blocking K channels in the cut axons, allows the Opto depolarization to be large enough to open voltage-gated Ca2_ channels and drive synaptic release. Thus, the cut axons as sufficiently hyperpolarized to sustain Na-channel dependent action potentials.

I think the best way for you to acquire knowledge and skill to understand optogenetics studies and to plan yours is to read recently published papers. First, you may need to focus your study on a specific function or dysfunction of the nervous system, and where in the brain you would like to see the effects of your experiments.

Then you can narrow down the papers that would help you to implement the right protocol for your stimulation and recordings.

Try the following to get a clue:

A pioneering paper on optogenetics by Prof. Deisseroth:

another one:

Development and application of optogenetics by Prof. Deisseroth team:

There are also occasional webinars related to optogenetics on the web.

Search for possible online courses too.

This vector from UPenn works well

AAV1.CaMKIIa.hChR2 (H134R)-mCherry.WPRE.hGH

Dear Kwanghoon,

I also can recommend the power meters and sensors recommended by Roni Hogri and Niraj S. Desai above (I find the S130C slim sensor quite useful).

However, these devices measure *power*, not intensity. In order to measure intensity with these, you need to know the total illuminated area on the sensor. That you can achieve in a number of ways, depending on what type of light source you are using.

1. If you have a uniform beam that has a larger diameter then the sensor aperture (9.5 mm diameter for the S130C), then you can estimate intensity as total power divided by sensor area (i.e. pi*r^2). Caveat: if you beam is not uniform, this estimate will be inaccurate.

2. If you have a uniform beam that is several mm in diameter, but difficult to measure the diameter exactly, you can place a smaller aperture onto the sensor, to limit how much light reaches the sensor. For example, if you drill a 3 mm diameter hole into piece of black material and place this over the sensor, and your beam is > 3 mm in diameter, then this would allow you to calculate the intensity as described in (1) above.

3. If you are illuminating a sample through a microscope objective, then these methods may not work well (especially if using a high magnification objective that typically generates a spot smaller than 1 mm). In that case, you would need to calculate (based on objective properties, wavelength, etc) the effective beam waist diameter at the focal plane. Use this together with the total power measured to obtain intensity.

4. If you are using a fiber to deliver the light, then all you need to know is the fiber core diameter (e.g. 0.2 mm) and the total power. As others have described, you just need to make sure that all of the light out of the fiber reaches the sensor, to get a measure of total power. Be careful because the fiber tip can scratch the sensor surface if they come in contact. In this case, you are estimating the intensity *at the tip* of the fiber. The intensity at the sample will most likely be much lower and generally quite difficult to estimate. But see:

That website is currently unavailable, so here's a snapshot from 6/2018:

For consistency, if you are doing in vivo experiments with an implantable fiber, I recommend measuring the total transmitted power *prior* to implanting the fiber "cannula", as this can vary from implant to implant (and hence animal to animal). You can then adjust the command power to achieve the same power delivered to the brain in each animal.

Thank you both for your help. It will help a lot

Hi David,

It's the photovoltaic effect, or the Becquerel effect (not the photoelectric effect). The photoelectric effect depends upon applied electrode voltage, the photovoltaic/Becquerel effect does not. I'm told there can also be an impact of the photoconductive effect, where light makes a material more conductive, which can also creates a voltage difference.

Anyway, the point is, it the effect occurs across the visible spectrum and down into the near IR.

It is a very common problem in optogenetic experiments. If you do a search you'll see lots of papers discussing it (e.g. Kozaia and Vazquez., 2015) . Theoretically, there are certain coatings that can be applied to electrodes (e.g. ITO) that will prevent the effect, but I don't think many people go to that extent.

If you're using MEA, you're actually quite lucky, as your light induced artifacts should be very reproducible from day to day, hence you can generate a recording of them with no cells on the array to generate a pure artifact, and then simply subtract this signal from your data.

> though it is absolutely suitable for most of the events in neurons, it may not be enough precise for ultra-fast source of current that changes in space within micro-second level

It is precise enough. Somata are isopotential on any biologically relevant timescape. You can model it yourself if you don't believe Rall and others. NEURON will happily split a soma into hundreds of compartments.

Hi Alberto,

Thank you for your response.

We have started adding HEPES to the incubating solution. The results are not very positive at this stage. See attached file, for the image of the STN in a slice that was incubated with HEPES solution.

Hi Ted,

Thank you for your response.

The neurons in other regions look better than the STN neurons and can be recorded from, unlike the STN neurons. See attached file for comparison between the SNc (substantia nigra pars compacta) and STN. It appears that STN neurons are perhaps more vulnerable to damage.

Thanks for sharing those references, we will take a look at them.

Interesting question.

I don't KNOW the answer. But I have a testable hypothesis.

My theory is that with optogenetic stimulation you are basically seeing temporal integration in SOM cells. That is to say, with your first light flash, you are causing say, 20% of SOM cells to spike, and the rest to be depolarized. On your second light flash, you are causing 40% of cells to spike, the increased number caused by residual depolarization left over from the first optogenetic exposure (i.e. the membrane is still charged up).

If this is the case, you should notice a change in the paired pulse ratio with varying light intensities, i.e. with very weak light intensities, you should see more facilitation than with very high intensities. Furthermore, the facilitation should not behave to changes in extracellular Ca2+ concentration in the expected way (though the first test is easier to perform), i.e. if you increase the extracellular Ca2+ concentration you should see no significant change in the paired pulse ratio.

Interested to see if anyone else has a better idea.

The first step is a light power meter, like a PM100D from Thorlabs with appropriate detector. Set that to the right wavelength, and read off the power in mW.

The you need to estimate the size of the excitation area. You probably have an etched calibration slide somewhere. Look down the scope, see what the diameter of your field of view is, and use 3.141*D to know your area.

Divide the first measurement by the area, and your done.

Yes, the measurement of your excitation area will not be 100% accurate, but it will be close enough.

Hello, have you successfully used collagenase to weaken the dura in the NHP?

Thank you, Thiago! That is what we need. I found on the internet "Databinge AAV Presentation" of Matthieu Vanni and Alexa Nelson. And using their recommendations I also chose UNC (it seems to be cheaper) and viruses AAV-hSyn-eArch3.0-EYFP or AAV-CAG-ArchT-tdTomado. Maybe Synapsin promoter is better (that to exclude the expression of rhodopsin in glia) but I heard that EYFP is not always visible. Thank you for your help. We will try AAV-hSyn-eArch3.0-EYFP virus.

Best regards,

Lena

Optogenetics is an emerging field that uses light-sensitive proteins to regulate biological activities in the body. The technique has been envisioned as a way to treat a range of diseases, including Parkinson’s and schizophrenia.

Optogenetics to precisely control cells to deliver insulin to diabetic patient. The approach could be used to continuously monitor blood glucose levels in human diabetics and automatically produce necessary insulin, a hormone that converts sugar from food into energy the body can use.

The researchers engineered human cells with a light-sensitive gene that is found in plants and produces insulin on cue when activated by wirelessly powered red LED lights. They inserted those lights and the designer cells onto small, flexible discs that were then grafted onto the backs of person or any place of the body.

The GIN mouse line (JAX 003718; Oliva Schwann JNeurosci 2000) exclusively labels Martinotti cells (MCs) in layer 5 of visual cortex, see Buchanan et al Neuron (2012) 75:451. It is a very sparse line, GAD67-based so post-hoc characterised as MC specific, it only gives you the tag, not the ChR2, and the genetic background might not be the best (albino), so perhaps not ideal for your purposes. To my knowledge, there is no Cre line specific for MCs. Any Sst-Cre line will drive at last three cell types, see McGarry Yuste Front Neural Circuits 2010, doi: 10.3389/fncir.2010.00012. I suggest the GIN line + patching + post-hoc verification with morphology and spiking pattern to achieve the best specificity for MCs.

Suggested experiments for determining if problem source is: primaries, secondaries, GFP cell marker misdirected to wrong cells or, virus. Primary Ab omissions are always useful to rule out/in some problems.

1) GFP primaries on sections from untreated wild type mice, secondaries as usual (2 slides, one rabbit and one chicken)...do you get label?

2) Primary omission on sections from untreated wild type mice, secondaries as usual (2 slides, one rabbit and one chicken)...if label above is it just due to secondary?

3) GFP primaries on sections from untreated (no virus) GFP mice, secondaries as usual (2 slides, Rb and Chicken).....glia and neurons?

4) Primary omission on sections from untreated (no virus) GFP mice, secondaries as usual (2 slides, Rb and Chicken ).....if neurons label in 3, is it secondary?

5) and 6).....as above 3 and 4 but GFP animals with virus injections (4 more slides total)

Notes: 1)use all fresh buffers and blocking solutions. 2)If slides are dear, just do 1-6 using only one or the other (Rb or chicken) primary Abs with its secondary Ab.

I am aware my comment may not be extremely useful as I am not familiar with the details of lasers used in 2P optogenetics, and not too familiar with the prices of Ti:Sa lasers.

However, I still want to ask: have you considered using a 1040nm laser to pump an optical parametric amplifier? Perhaps such a setup would still be within the budget? 920nm is a wavelength easily accessible for an OPA pumped with the SH of 1040nm lasers.

If you are using a laser with near-infrared emission, you don't need to warry about the damage problem at the output end facet. However, at the input facet I suggest you use a fused splicing method, rather than direct  mechanical connecting or too tight a focusing. If you want to launch a free-space beam into a fiber patch cord, you should use a NA-adapted focusing component.

Take a look at this lab: http://denizatasoylab.com/

 Hi Ariadne,

I think that counting fluorescent cells and dividing by fluorescent cells plus non- fluorescent cells is pretty standard. Neurons are pretty easy to pick it in culture, so unusually, people just pick a few fields of view to make the quantitation.

FACS is more accurate, but then you have to worry about glia. You might try NeuN staining to isolate the neurons, though, if you decide to go that route. 

Best of luck, 

Joey 

 In order to find out whether Refik is right and it is an "electrical artifact", did you trigger the laser, but had unplugged the fiber before, so no light reaches the tissue? And I really mean UNPLUGGED aka physicallyY BLOCK the LIGHT?  If you still see an artifact, the method of Refik might work... or some more thoughts about ground connections in your setup. 

If it really is the light itself - is it intensity dependent? If you need to stick with single wires, go for stainless steel or Pt (FHC and many others...) or glass coated PtIr  multisites (AlphaOmega or Thomas Recording). We do see light artifacts when illuminating MEA arrays - but I doubt it is a very specific photoelectric effect...  More of a thermal-electric effect (thats why I was asking about the light intensity dependence...)  

If you go to apply silicon probes, be aware, that (very much dependent on the lithography process), the interface between metal and electrolyte might affect the semiconductor below. It may very well be a depletion layer down there, thus establishing a real photodiode perpendicular to the electrode into the Si-bulk.... Thats why I prefer Si-probes based on the Silicon-on-Insulator over p-doped Si-probes.... ;-)  So, I am not quite sure which process Neuronexus is using for all their probes (it is recognizable from their spec sheet), but you might want to look into the Atlas Si-Probes as alternative... Their newest ones should not show these photo-currents. 

And last but not least, refering to template-subtraction: An artifact is an artifact and if it is big enough to be a nuisance, it's subtraction is big enough to spoil your data. 

I know everybody is using this method, but I am not really fond of it - but alternatively blanking your signal is really worse!

Good luck!

uli

Hi 

I would suggest you to use DAT-cre mice since TH is not exclusively expressed in DA neurons. 

There have been several studies using transgenic mouse lines, injecting with viruses in VTA and then stimulate with optical fibres the terminal areas to study behavior. You can have a look on those for example:

https://www.ncbi.nlm.nih.gov/pubmed/?term=steinberg+plos+one+nucleus+accumbens (TH-Cre rats though) 

https://www.ncbi.nlm.nih.gov/pubmed/?term=berrios+nature+communications+2016  (TH-Cre mice) 

https://www.ncbi.nlm.nih.gov/pubmed/22445345 (VGAT-cre mice)

https://www.ncbi.nlm.nih.gov/pubmed/?term=qi+morales+nature+neuroscience+2016 (Vglut2 Cre mice)

Good luck!

Zisis

The ventral vs dorsal dichotomy originates from some differences in object processing correlating with the lesion and functional activation studies, e.g. prosopagnosia with right  temporobasal lesion / activation focus and pure alexia in the homologue regions in the left. Positional tasks and spatial tasks comprising tasks with respect to the own body are correlated to dorsal lesions / activations. From a clinical / lesional pov, already the cerebral akinetopsia (lesion at the lateral temporooccipital junction) raise some concern. In my opinion, connections from primary brain areas in a modality to other modalities should be more accounted for, and of course there must be a connection between dorsal and ventral pathways, which makes this dichotomy a good similar kick-off to test hypothesis' against as it was the case in language localisation.

Have some idea from here too..

Hi Chelsie,

I have no experience investigating fly vision or long-term red light exposure, but I have used CsChrimson extensively over the past 2 years, both in explant brains (preferably) and in vivo, and might have some clues to what you're seeing.

First of all, my colleagues and I all see some Chrimson activation also in non-retinal fed controls, presumably because the intrinsic levels of retinal in the fly are sufficient to drive Chrimson to some degree. Looking through my data, the non-retinal controls are sometimes responding up to about 50% of those fed retinal (this is GCaMP6f response of downstream neurons).

Secondly, flies do see the red light (I use 630nm). It is sometimes suggested that flies can't see far-red light, presumably because the rhodopsin sensitivity doesn't seem to reach much further than 600nm (see Salcedo et al., 1999, JNeurosci), but my colleagues and I have indeed found subtle behavioural as well as neural responses to light >600nm. Presumably Rh6 still has sufficient sensitivity at 630nm if exposed to light with high enough intensity.

So, to sum up, your effects could come from residual Chrimson activity in your controls, or from the light itself. Have you already tested the genetic controls not expressing Chrimson (or any other red-shifted channelrhodopsin, for that matter)?

Cheers,

Oliver

rAAV2-retro-Syn-Cre into the target area, then inject flex-[favourite inhibitory opsin] into your secondary area seems like a good place to start. So long as your secondary area is a good few millimeters away, you shouldn't get direct diffusion of the rAAV2-retro, and instead would rely on retrograde transport to bring it to where your flex-inhibitory opsin is.

I use [favourite inhibitory opsin] as opposed to recommending a specific opsin, as I've found both eNpHR3.0 and eArchT3.0 a bit unreliable at shutting down neurons completely. 

Hi,

Please read this article it sounds they covered different protocols for  activation of channelrhodopsin, however its in nervous cells, but you may optimise the protocol yourself to suit your experiment conditions  

Hi,

Unfortunately nobody answered, I thought you can help me. 

Furthermore Kendall is not very helpful. I asked them, but they did not reply. Today I will Skype with another company, Cambridge Neurotech. In next three month I have to buy some reliable, radio wave guided system, or I lost my grant money.

If you  have any information or suggestion, please share with me.

Bests, Kornél

Thank you everyone for your advice.

As an update, I just got the histology on another animal in which I injected three distinct vectors in three regions, one being my vector that did not show expression previously.  I injected the problem vector into the sensory cortex this time, and there was still no expression.

I did get expression in the BLA with a different vector (same serotype, same promoter).

Therefore, I believe it is the vector itself that was ineffective, even though the reported titer was high (6 x 10^12 vg/mL).

Hi Cassey,

the short answer is yes, AAVs are able to infect axon terminals and produce retrograde transport (towards the cell body). The slightly longer answer is that this process is highly biased based on the seropype you are using. There are a number of papers reporting AAV6 (and maybe AAV6.2) is the main serotype to produce retrograde transport. There are however reports of many other serotypes producing retrograde transport with a much smaller rate. Keep in mind that there are many other considerations related to your experimental design that will help you minimize the consequences of this possible off target effect. One of the most obvious, since you talk about a cre line, is that if the axons that get infected are from a Cre- cell then the expression of your gene of interest won't take place; for example if you use a Dat Cre of a Gad Cre mouse line, terminals from pyramidal cells could be infected but they won't be able to express. Moreover, if you are interested in activate a specific cell population in many cases you apply the light stimulation (in the case of optogenetics) in the area where the neurons you infect project, and not in the area of injection itself since this approach increases the specificity of the manipulation. Finally retrograde transport, with AAV, HSV or CAV2, can be used to your advantage as it allows you to be much more selective in your manipulations. There are numerous recent papers taking advantage of this approach.

Hope this helps,

Ezequiel

Hi Miguel,

Assuming you are talking about AAVs, we have very good results regarding viral spread and expression in cortical neurons with AAV9. AAV5 also works very well. Even though there are publications showing infection and expression with AAV2 we never had god success with that serotype so I would stay away from it.

Hope this helps,

Ezequiel

Hi Alessandra, Thank you for the information!  The difference though is that my experiments are un-conditioned real time place preference...It is a novel exposure experiment with one side paired with optogenetic light activation.  At any rate, these are good resources for me to know in the future if I plan a CPP test! Cheers,

Leandra

Hi Tsai, I hope you realise this video you linked is a master thesis but not for science but science communication... 

Neuronexus has commercially available optrodes which can be used chronically in freely moving animals. 

In addition, the opeOptogenetics community has these approaches collated:

I hope this helps,

Ede

there should be no problem using the same fiber.

higher NA is not needed

In injecting the oxytocin neurons, where did you inject?  Not all OT neurons are similar in terms of the connectivity.

If you pull your fiber too far from your target, you may get false negatives in your behavioral outcome...

Alberto makes an important point.  The seizures could be less a function of your level of stimulation and more related to the brain region where ChR2 is expressed. If you have intentionally or unintentionally included the endopiriform (subregion of the piriform cortex and closely connected to entorhinal) into your list of infected regions, then you could certainly stimulate seizures, as it is a source for seizure activity (see: http://jn.physiology.org/content/86/5/2445.long).  Since you are seeing this even at lower power ranges it starts to rule out activation intensity as a cause.  20mW is a lot of power indeed, but other targeted regions have been published to require 20-50mW as a minimum level of stimulation needed to attain their behavioral effects.  

The endopiriform (in mouse/rat) lies just lateral to the external capsule (see attachment), and that capsule (,"ec" purple arrowhead) acts like a liquid channel, which will conduct your viral injection away from your desired target right down to the endopiriform (black arrow) and possibly the lateral amygdala.  So there could have been unintentional infection of this epicenter of epileptiform activity.  Even if you were mircoinjecting distant structures like the dorsal hippocampus, you can still wind up with virus in the endopiriform with the slightest excess or spillover (by the way, in case you aren't doing this already, be sure to wait 5min after injection is complete before raising your needle, and even then only raise it another 0.2-0.3mm and leave it again for an additional 5min before completely removing your needle to create an air pocket and allow absorption of viral suspension and avoid spillover that can occur with premature removal of your needle).  It would be best to check reporter histology in this area to first confirm/deny this.  

This potential cause I described is all less likely of course, if your target region is much further away (>1mm) as the others have described here.

the pulses are defined in miliseconds, rather than HZ.

in order to program a 50 hz pulse, for instance, you need program a 20ms interval and a 20ms pulse duration.

the answer by refik is pretty accurate.

if you need pulses with decimal point timing (2.5ms, for instance) please contact PRIZMATIX support and we will provide you with a software update.

arie

A little late to the party, but we use AnyMaze for Opto+Maze tracking simultaneously. If you still have questions let me know and I will do my best to answer. The system works well and the optic fibers rarely cause any issues with tracking.

Hello Shan,

I am using the UHP-Mic-LED-460 from the same company to stimulate my sample during electrophysiological recordings. At first I had some noise from the source, but then when I isolated all the wires that necessarily go into the faraday cage (simply by covering them with aluminium foil and then grounded) the noise disappeared completely. So, in my experience it is not particularly noisy,

good luck

Hi, the linked ones are reliable manufacturers, but there must be many other. And these lasers can be suitable for you.

If I'm right you measure the laser power after the fiber. Is there any adjustment possibility at the fiber coupling? Its possible that your laser is stil perfect just the coupling needs to be fine tuned.

Regarding your post-laser LFP observations:

I would expect the broad silencing of an entire population of important inhibitory neurons in V1, such as you describe here, to rapidly generate massive excitation across most or all layers. Due to recurrent connections across layers and within layers, and depending somewhat on the duration of the laser exposure, perhaps the PV-silencing leads to local seizures.

Depending on the duration of the laser exposure, and how many times this was repeated, it is possible that repeated, prolonged light-induced seizures have resulted in cell death and/or detrimental plasticity, essentially making V1 epileptic. Then what you are seeing in the post-laser LFP could be "spontaneous", local seizures.

Regarding the "pre-Laser" differences in LFP with respect to other mouse lines:

I wonder whether - depending on expression level and method - there might be some low-level activation (and hence proton-pumping) occurring even without exposure to green light, or perhaps at ambient light levels? At first this seems unlikely, but even a very small effect would have time to accumulate over weeks. But in any case you could compare whole-cell recordings from ArchT-expressing neurons and PV+ neurons in wild type, or by crossing PV-Cre with e.g. a tdTomato reporter line, and look for any differences in intrinsic properties (such as greater excitability, lower AP threshold in the ArchT mice).

It sounds like the CAV-Cre + AAV-DIO-opsin virus combo would be best.

I've used WGA alone as a tracer in the brain and via virus as AAV-WGA-Cre, and it kind of goes everywhere. WGA is quickly transported transsynaptically retrogradely and anterogradely through most neural circuits. Thus, the issue that occurs is whether the DIO construct is being expressed in first, second, or even third order neurons in relation to the AAV-WGA-Cre injection site. For this reason I'd avoid AAV-WGA-Cre, regardless of serotype.

Replication-incompetent PRV would work better as it would go monosynaptically retrogradely (depending on serotype, I think), but it requires biosafety level 2 facilities and quarantine protocols. I also haven't heard of labs that distribute PRV-Cre viruses, though they may exist.

The CAV virus infects cells monosynaptically retrogradely, it only requires BSL1 facilities, and there is a distributor for it in France. I personally haven't used it but I know people at my university that have used it in combination with a Cre-dependent AAV construct (DIO-DREADD instead of opsin) with reasonable success. The timeline is the same as a non-Cre dependent AAV for optimal opsin expression, about 4-6 weeks post-injection.

Good luck!