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Stereotactic Surgery - Science topic

Stereotactic surgery or stereotaxy (not to be confused with the virtuality concept of stereotaxy) is a minimally invasive form of surgical intervention which makes use of a three-dimensional coordinate system to locate small targets inside the body and to perform on them some action such as ablation, biopsy, lesion, injection, stimulation, implantation, radiosurgery (SRS) etc.
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I need to make two consecutive stereotactic injections in the same area of the midbrain of a mouse. How can I do this correctly without traumatizing the mice? Are there tools to maintain drilled holes in the mouse skull for repeated injections?
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What time period is between these injections?
Have you considered inserting a cannula? this will allow for repeated injections into the brain.
You could even insert an osmotic pump if you wanted continuous infusion. this is less traumatic than even the cannula, and will require less equipment.
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I will attempt to keep this as concise as possible, as this has been a toilsome endeavor.
1st issue:
Our coordinates never exactly match up with what the brain atlas says. The virus I inject almost always appears to be more dorsal than intended. It is to the extent that I have aimed towards the bottom of the basolateral amygdala and I have barely reached the top of the BLA. I purposefully picked -1.3mm for the anterior/posterior coordinates and +- 3.0 mm for the medial/lateral coordinates as this part of the brain atlas showed smaller fiber tracts to make it through and it was easier to make it around the ventricle. This also gets me some wiggle room as the BLA is fairly large here and neighboring anterior/posterior coordinates.
2nd issue:
We also seem to have sporadic unilateral injections even though we are doing bilateral injection.
I draw up 950nl into a Hamilton syringe. I eject 50 nl to check virus has been drawn up by the virus. After injecting over 5 minutes, I let it diffuse for 10 minutes. Then I pull the needle out and eject 50 nl to ensure that the needle wasn't clogged. I do my 2nd injection. Repeat injection and diffusion times. Eject 50 nl and check to ensure it wasn't clogged. So I dont understand, how it is even possible to have a unilateral injection.....unless it is getting caught in a ventricle?
If anyone has any thoughts about what it could be or even thoughts about how I could better troubleshoot, please let me know.
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Hi Mark,
Accurate targeting is always an issue for stereotaxic surgery! I do not have much experience in terms of injecting BLA. Here are some general thoughts about your issues.
1st issue: Brain atlas were mapped using a specific age animals (also within certain body weight range). If the animals you used are a little bit far from that range, you should adjust the brain atlas coordinates by testing a few injections. For details, if you have dorsal-ventral targeting issue, first make sure the animal's skull are leveled by making the bregma and lambda on the same horizontal level ( I guess you already know this.). Second, different people may have different habit to define the top of the skull where you set as 0 point for your dorsal-ventral injection. If you keep injecting too dorsal, you can define your top of the skull a little bit deeper.
2nd issue: If you do not see injected virus, there are a few possible reasons. You already mentioned some. First, the needle could be blocked. Second, the virus could be injected to ventricle. However, it could also be due to the way you examine the virus expression. One possible scenario: if your needle had some bending during the injection, it may cause some off-targeting in the anterior-posterior axis. And if you only check the BLA region, you may miss the injected virus in the nearby regions in terms of anterior-posterior axis.
Hope these thoughts could help you solve your problems.
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We tried different methods. The problem consists of the absorption of our colour marker by the brain tissue. We want to reach to the basal forebrain, 5.5 mm deep from above by cranial stereotactic surgery with canulas of 0.43 mm diameter. For those trials we kill the mice and do the surgery post mortem.
Coating the glass fibre canulas with ink or rhodamine solution didn't work so far, it seems to have been wiped off while entering the brain tissue. We cannot see the hole/tunnel made by the glass fiber canula, probably because the brain tissue wobbles back into place after ejection of the canula.
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Maxim Telle, in this case, the over warming of the tip of the canulas needs to produce perceivable local edema.
The over warming has to discernably increase the osmotic tissue pressure at the canulas' tip.
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I recently started learning stereotaxic injections in mice, we will be targeting the Medulla but I started with practicing ICV to get used to the process using Evan's Blue Dye 1:100, and I had a very few success and too many failures. I don't have trouble with fixating the head as when I started, and I think I don't have trouble reading the coordinates measurements in the Vernier scale.
The coordinates we use for right ICV injection are, from the Bregma:
  • Lateral (X) = -0.9 mm
  • Posterior (Y) = -0.1 mm
  • Ventral (Z) = -3.1 mm.
Most of the time I get results as below, what might be the reason?
Thanks a lot in advance.
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Dear Sir Al-Hamamy,
I see that you have posed several matters in the same topic. They must be spliced to explain better.
1 - The dye do not spread to the contralateral ventricle
ANSWER: This not probable to occur, unless you inject very large amounts of dye. This is not recommended why this would injury the animal. Even more, the normal flow of the cerebrospinal fluid tends to carry the dye rather and firstly to the 4th ventricle (that is a median cleft in the diencephalon). If you aim both ventricles, you must perform two separate injection in two separate coordinates.
2 - You target the medulla? Which structure specifically?
ANSWER: You must be very careful why the medulla contains the pre-Botzinger complex, that is the primary respiratory drive. This is complicated by the very small volume of the medulla, which is not as complacent as the telencephalon structures. So, injections of very small amounts, even less than one microliter, in this region can cause terminal apnoea. Also, to target specific nuclei in the medulla, you probable will need a stereotaxic micromanipulator with a precision of 1/100 micrometer or greater instead of the more widely available micromanipulators which have a precision of 1/10 micrometer.
3 - Please, do not begin the surgery before you are absolutely sure that the head is properly fixed and centered in the holder. You must begin putting the animal's incisor teeth in the incisor bar. After that, you must palpate the ear meatus and plug it with the ear bar. It is a little difficult for beginners. The gold standard is the blinking of the same side eye due to the compression of the facial nerve towards the border of the ear meatus. Once both meatuses are plugged with the ear bars, you must center the animal's head using the ear bar verniers.
4 - You must be skilled in vernier reading. You can train with a calliper.
5 - I strongly recommend you to read:
- FERRY, GERVASONI, VOGT 2014 - Stereotaxic Neurosurgery in Laboratory Rodent
- The preface of the Paxino's atlas, where the authors teach how to deal with craniometric points and coordinates.
I hope I have helped you.
Sincerely
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Hi, I am Joe and currently, I am working in a neuroscience lab using the optogenetic technique. And I have some technical problem with that, which the AAV always injected off target, for example when I inject the AAV in the BLA. However, there is always a lack to the CeA and the injected passage which bothers me a lot.
Therefore I would like to ask if there are any tips to prevent the situation that I mentioned? Thank you so much for your help.
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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.
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Surgical lesioning, like thalamotomy, is usually targeted using AC-PC coordinates. Does anyone know of thalamotomy coordinates in MNI space?
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Bassam Al-Fatly there is a nice online tool which can convert Talairch (AC-centered) coordinates to its MNI equivalent. It can be found here: https://bioimagesuiteweb.github.io/webapp/mni2tal.html
If you don't wish to convert and simply want a commonly used probabilistic MNI ViM location, this paper lists X=13.05, Y= -18.36, Z = -2.01 as a good coordinate to use to target this thalamic nucleus :
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Hi All,
I have been running into a technical issue when I try to cannulate the prelimbic cortex of mice. I find that I am about .1-.2 off from being even (see the attached pictures). I am using bilateral cannula with .8 mm between the tubing. I found coordinates (AP: 1.78, ML: ± 0.4, DV: -1.5) that get me to the right place A/P-wise but my problem is M/L. I am careful with how I place the mouse in the stereotaxic frame and I check bregma and lambda prior to drilling my screw and cannula holes to make sure the brain is level. I suspect I am setting my reference (i.e. bregma) incorrectly but I don't see how. Has anyone experienced this and if so, how did it you fix it?
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What are you using to hold you r bilateral cannula during placement? It appears to me that your cannula is not level when it is placed. One way to check this would be to make sure the Dorsal/Ventral(DV) coordinates are exactly the same on both sides when the cannula touches the skull before drilling holes. If they are off of level by even tenths of a millimeter it can make a HUGE difference in placement. I would also caution you to wait a significant amount of time after dental glue placement, before removing said cannula from the holder as any slight movements before the glue is completely set will cause change in placement.
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Hi, I am currently learning to do electrophysiological recording on mice visual cortex, and sometimes the cortex bleeds and swells after the craniotomy and removal of the meninges. I want to know if bleeding is the major cause of the swelling cortex, and how to prevent the cortex from swelling.
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Dear Zhou,
It is important not to cause any damage to the brain tissue-paranchyma at any stage of surgery. First remove the overlaying skin, then the bone carefully with care not too cause damage to the underlying membranes or excessive bleeding that may increse the pressure. Before cutting duramater, make sure all the bleeding is stopped- if the bleeding is coming from the bone, use wax to block it. A cautorizer with a small tip and surgucell may help with soft tissue bleeding. Once you remove the duramater, you will get to the piamater with many fine vessels and capillaries, make sure you do not touch or pinch this membrane. From now on you will have to keep the brain tissue moist either by warmed up saline or inert oil such as silicon oil. Another point is to use/make electrodes with long and thin shafts, so that the tissue damage due to multiple electrode penetrations will be limited-otherwise large diameter electrodes will make a large hole-damage on the surface, especially if you need to go deep into the brain.
Please have a look at my earlier studies and thesis for details of steretaxic surgery and methodology.
Best wishes.
Refik
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Hello, I'm recording neurons in vivo in the ventral tegmental area of rat brains using stereotaxic equipment. What is an effective way of determining whether my equipment is accurately horizontal and/or perpendicular with relation to the animal other than just "eye-balling" it? The VTA is 7-8 mm deep and I'm attempting to target an area that is roughly 0.4mm wide and 0.8 mm long so even small deviations from being being perfectly perpendicular will have an effect.
A related question: how can I ensure that the head of the animal is properly secured in a position that is perfectly horizontal?
Any help is greatly appreciated. Thank you.
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Alignment indicator is a nice thing, but costs several thousand $$. We usually spent a lot of time leveling the skull horizontally until we developed a simple bubble (spirit) level probe ( http://www.invilog.com/products-services/auxiliary-equipment/making-sure-the-skull-is-in-a-level-position-takes-a-mere-2-minutes-with-the-invilog-bubble-level-probe/ ) which is working either in rats or in mice.
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Hi everyone, 
I am performing stereotactic surgery for mPFC in CD1 mice. The coordinates that I am trying to use is AP: +1.9, ML: +/- 0.3, DV: -2.5. Apart from this antero-posterior coordinates, I have also tried +1.6, +1.7 and +1.8. But in all these cases, I see enormous blood while inserting the cannula or even the injector. I see so many papers injecting drugs, lentivirus etc in mPFC using same coordinates, but none say a word about the blood. If anyone has experience on this, please share your experience here. Thanks! 
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This is unfortunately a very common problem. There is a few things you can do. Rather than using a drill to make a hole to insert a cannula or other instrument, I simply start the hole with the drill and then finish it with a 27g needle. I find that making the hole manually is more delicate and it gives you greater control over adjusting before you end up with a massive bleed. Depending on what you are doing, another tactic that people also use is targeting at an angle to avoid the problem completely. I have seen this done in papers, but have not tried it personally. I think this would be difficult with an infusion needle unless it was very stiff, and you'd have to practice adjusting your equipment to be able to insert at an angle. 
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Instrument would be used for experimentation in mice.
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I am trying to find a reference describing the accurate method to locate the lateral ventricles for a female Sprague dawley rats using stereotaxic apparatus.
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Dear Beatriz
If we can use the same co ordinates for animals upto 400  g, why the atlus is strict to use coordinates  upto 250 g?
Thanks
Smriti
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The setup is isoflurane with passive scavenging through an activated charcoal filter. These mice are Parvalbumin Cre (+) and (-). Mice are placed on a heating pad throughout surgery. There also is a lamp about a foot and a half above the surgery table that can get pretty hot. I'm not sure if this is the problem though. The mice have no problem going down. Their breathing is normal up until about an hour to an hour and a half. Their breathing becomes labored, they start pooping, and their muscles become rigid. Although their breathing gets lighter, they do not respond to a tail or toe pinch reflex. After about 5-10 minutes of this labored breathing, they die. The isoflurane level is between 0.5 and 2 during surgery.
Do you think it is a problem with the isoflurane chamber, hyperthermia, buildup up isoflurane waste, or something else? Any help would be greatly appreciated. I have attached a video of the labored breathing with the mouse not responding to a touch pinch.
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Dear Sam,
We've been doing quite a lot of stereotaxic surgery usually administering AAV's into the hypothalamus and amygdala and it is always very important to determine that both your skills and set-up are appropriate. Can I ask if there are other users that successfully use your set-up? If so what do they do differently? Are your surgeries longer than others performing a similar protocol? If you are just learning the procedure nothing beats watching someone skilled in the procedure to pick-up any necessary tips from them.
In the past, we had a few problems with our initial anesthetic set-up using Isoflurane, it just needed some slight modifications. This was largely due to either providing too much isoflurane or having the scavenger set at too high a level. You can tell if the mouse is getting too much anesthetic - their breathing will get quite labored and they'll start to gasp. If they are getting too little then they'll start to squirm as they gain consciousness. Remember that the scavenger canister needs to be changed regularly and if you have a new canister it will scavenger much more than one that is near-full. Isoflurane is a good choice of anesthetic but needs careful monitoring throughout the surgery. It really shouldn't need to be adjusted throughout. We normally have it set around 1.5. I'd think that if you are using less than 1 then something is not set-up correctly. If the animals have slow steady breathing then you know that the anaesthetic set-up is appropriate. 
In regards to temperature, if you are using a homeothermic blanket then the temperature should not be an issue as the blanket will regulate it appropriately. If you don't have a homeothermic blanket I'd suggest that you get one. Heating pads/blankets are alright for post-surgery recovery but shouldn't be used during the surgery itself.
Can I also ask if you are using any analgesic? If so, is the dose appropriate? If the animals are dying after about an hour, it might be an overdose from the analgesic.
I agree that you should perform a test to determine if the surgery is killing the animal. This could be done just placing a mouse in the frame as usual and monitoring it as you would during a surgery for the same amount of time. If the mouse survives than you know it's the surgery and not the set-up and can seek some expertise on the surgery itself.
Hope this helps. 
J
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I am injecting 200nL of a vasoconstrictor into mouse brains, trying to hit a very precise area.  Can I inject a small amount of india ink with my vasoconstrictor as a way to see post-tissue processing if the injection is exactly where I need it?  I am doing survival up to 3 weeks, so would india ink be stable and not interfere with flourescent IHC over that time frame? 
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I would recommend you use the damage to the area as a marker. Damaged tissue will autofluoresce in all channels and this is easy to see even months later. I was able to verify hundreds of injections this way. I use a 33G needle and I inject 5mm deep, but the entire needle track, including the tip, is easily visible under fluorescence. In fact I have to image an extra channel in order to subtract out the damage. I can send pictures if you want.
If you are using a cannula, you can inject trypan blue or whatever just after perfusion, which is what I do for cannulated animals. Otherwise I inject a fluorescent marker, like CTB or fluorogold, but as stated before, this may interfere with your actual experiment, and you would have to ensure that it does not before proceeding.
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Has anyone performed stereotaxic surgeries into the BLA from an angle rather than straight down? I would like to angle the injection so that i do not damage the striatum, but don't know a good angle to approach the BLA. 
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Thank you. Do you have a protocol you could use? thanks again 
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I need to make sure whether I did correctly in the surgery. I need to implant tetrodes in the mPFC of 4 month old rats. I have a confusion about dura removal. After removing a small piece of dura, I could see the thick white layer shown in the attached file. And when I removed the white layer, I could see clearly the vessels. Here comes the problem. Sometimes, even after removing the white layer, my tetrodes still couldn't get through. I have to remove another transparent thin layer containing vessels. But sometimes, my tetrodes could get through without removing the transparent vessel layer. So I am thinking whether below the white thick layer, there are acturally two transparent layers. For tetrodes implantation, I need to remove the first transparent layer and leave the second transparent vessel layer?  But in my operation, sometimes I remove both transparent layers and cause bleeding.  I searched the structure of meninges, and I got to know dura consisted of two parts. So I am curious the white thick layer is only one part of dura or what? And what about the arachnoid and pia mater? I couldn't see them under microscope and I don't need to remove them for my tetrodes, right?
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 Hi Tianyang,
A picture says a thousand words. I didn't really get your question, but actually a really easy way to know that you haven't removed dura, is by seeing how the light shines off the tissue, if its pretty reflective, you probably haven't removed dura. In this pic I would say the dura is still on (though it is a bit out of focus).
Here is what to do, take a very fine needle (30 gauge) and slightly bend the tip of the needle so it resembles a hook of sorts. Then under the scope, poke around outside your area of interest (obviously carefully). You should be able to stick the point through the dura (and maybe a bit of cortex) and then lift the dura so it separates from the brain tissue. That is how you will know what to remove. From there you can use the ripped dura as an entry point to start cutting dura 2. Bleeding; blood vessles are every surgeons' nightmare, so you are not alone. Just try to avoid breaking it when you remove the dura, sometimes you can't avoid it. Have a syringe with saline prepared in a bucket of ice, and gently spray the area with saline until it stops bleeding (I also recommend non-toxic absorbant gelfoam). Good luck! 
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I am seeking a protocol or advice on expression in vivo. I am currently using a GEG Tech lentiviral vector containting GFP and a NMDAR subunit gene. I have a physical titer of 4.5E6 RNA genome/uL and an efficient titer of 1.9 E9 Transduction Units/mL. I have not been able to detect any GFP expression, and would like to rule out delivery as the problem. I have been experimenting with volumes ranging from .5, 1, 1.5, 2ul per injection site. Injections are done manually using a micropipette with a manual delivery system. I typically allow 5-10 minutes for viral absorbtion before removing the needle. In histology of the injection site, I see granules that fluoresce in all spectra, but these do not seem to be cells. Any advice or wisdom about lentiviral injection is appreciated.
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Some colleagues of mine just published a nice methods paper related to viral vector injections:
Curr Protoc Neurosci. 2015 Oct 1;73:4.37.1-4.37.31. doi: 10.1002/0471142301.ns0437s73.
Use of Adeno-Associated and Herpes Simplex Viral Vectors for In Vivo Neuronal Expression in Mice.
Penrod RD1, Wells AM1, Carlezon WA Jr1, Cowan CW1.
Author information
 
Abstract
Adeno-associated viruses and the herpes simplex virus are the two most widely used vectors for the in vivo expression of exogenous genes. Advances in the development of these vectors have enabled remarkable temporal and spatial control of gene expression. This unit provides methods for storing, delivering, and verifying expression of adeno-associated and herpes simplex viruses in the adult mouse brain. It also describes important considerations for experiments using in vivo expression of these viral vectors, including serotype and promoter selection, as well as timing of expression. Additional protocols are provided that describe methods for preliminary experiments to determine the appropriate conditions for in vivo delivery. © 2015 by John Wiley & Sons, Inc.
Copyright © 2015 John Wiley & Sons, Inc.
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For example, if I inject 1 uL of drug solution directly to the primary visual cortex, what is the dispersal volume for which I can expect a drug effect?
1 mm^3 ?  
5 mm^3 ?
References appreciated.
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In this study, radio-labeled dopamine was injected intracerebral (IC) at various volumes into the brainstem and then radioactivity was measured from punches in series of coronal sections at two different time points (1 and 15 min post-injection). Then they plotted the proportion of radioactivity in the lateral, horizontal, and anterior-posterior planes, relative to the injection site. 
Gives a sense of dispersion - and was helpful.
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I am injecting AAV cre into the dorsal hippocampus in 4-5 week old mice. The coordinates I have are -1.5mm [AP], 1mm [lat], 1.5 mm [ventral]. However these coordinates are for adult mice, and I want to inject in 4-5 week old mice. I'd like a reference point to where I should first try injecting for the dorsal hippocampus. 
It would help me if there was a paper that did stereotaxic injections in young, and adult mice and what the coordinates were for those two mice. I had a difficult time finding a paper that had this.  
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Hi Thomas,
I am agree too. You need to adjust your coordinates. I calculate a ratio. I use C57BL/6J mice and the published distance between bregma and lambda for these animals is 4:21mm (3 moth old around 26-30g). If your animals are smaller, the distance between bregma and lambda is going to change. Then you take the distance between bregma and lambda of your animal and divided by 4.21mm. This value then has to be multiplied by your coordinates. Now you have stereotaxic coordinates adjusted for the size of your mouse brain.
As a reference I used Moore and Boehm, 2009.
I hope that this help.
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Need to inject ASOs into 4 weeks mice. Any suggestions?
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Which ventricle are you going into? Will you do a single injection or implant a cannula?
LV is quite large and easy to target. I would start with +/- 1.0, -0.2, -2.0 (x,y,z) and do some test runs with trypan blue or another dye. If you see dye in the 3V then your coordinates are good. These coordinates are just a starting point, and calculated for adult mice. You should refine them for your mice and your stereotax. You may also need to adjust for younger mice.
3V is much more difficult to target because it's deeper and very thin. I would start with 0.0, -1.8, -5.3 (x,y,z) and do test runs with dye as above.
If you are implanting a cannula, a good way to check the placement is if you see a small droplet of CSF when you remove the plug (dummy cannula). If you implant at 4w and then let them age, the cannula may move out of alignment as the brain grows.
Another thing to keep in mind is that inbred strains like B6 and 129 have a ~15 degree slant to their skull relative to the brain atlas, so I find that I have to adjust my coordinates to be more anterior when injecting deeper regions.
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I am working on an endogenous brain tumor model and will induce tumor formation by stereotaxic injection.
Part of the treatment regimen will also be delivered via stereotaxic injection into the tumor tissues - is it possible to perform this procedure twice in the same area within 2-6 weeks? 
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Yes. I have done that. Rabies tracing experiments require 2 injections about 3 weeks apart.
Here is a reference where they do it. (Fig. 1B)
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I am planning to perform a pharmacological study to assess hippocampal neurogenesis via IGF-1 receptor inhibitor. But most studies were performed  in vitro or in vivo (systemic). So whether this inhibitor of IGF-1R (such as AG-1024 or picropodophyllin) can be  intracranially injected by stereotaxic localization and what is the optimal dose? Thank you!
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Ideally, you can infuse over hippocampus, because as mentioned by Giovanna you can induce gliosis in hippocampus.
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Thanks in advance for your replies.
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We use blunt earbars that do not penetrate the eardrumm. Kopf model nr 855, 955 or 1755. Specs: 45° Non-Rupture tip with a 0.8 mm radius.
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I will use it for functional and stereotactic surgery planning.
I use a CRW frame with which I use a very simple software for planning, and I am in need of a more sophisticated and low priced software.
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Take a look at Osirix. There is a free open source version with many plug-ins available. I have not used it for stereotactic planning, but I believe this functionality is available.
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I'm trying to find a device to be attached to my stereotactic to perform bilateral injections at once.
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Thanks Ulrich for your response. Unfortunately,  I'm not very good at building homemade devices. Even less if it has to be precise enough to inject something in delimited brain nuclei.
Regarding the two-arm stereotax. Maybe it could do the job. However, as far as I know, I would have to manage each arm separately, isn't it?. At the moment, I'm doing one injection in one nucleus and then moving to the other lateral (using a conventional one-arm stereotaxic instrument). So, I do not see a great advantage using the two-arm stereotaxic. However, I may be wrong because I have not much experience with two-arm stereotax.
Thanks again for your ideas. 
Juan
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Our lab is about to purchase several new microscopes for implantations and microinjections to mouse brain, and we've never really been thrilled with the scopes we've had at our disposal. I'd appreciate if anyone has recommendations of scopes they enjoy using.
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We do not use a microscope anymore - instead we equipped a USB-CCD with a "telescope lens" on a Manfrotto-arm (or the like) and display the surgical area on the screen. Great for screen shots.
Something like this: "imaging zoom lens mounted to a CCD camera on a stand, which was independent from the stereotaxic frame (Imaging Zoom Lens, 0.7X-4.5X, NT53-347, Steel post components NT03-609, NT58- 994, NT58-955, NT58-974, NT54-976, Edmund Optics, FireWire CCD Color Camera DFK 41BF02.H, The Imaging Source)" (from the Thesis of Susanne Löffler)
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We are developing such system and we are intersted in similar developments.
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What do you mean by "completely automated stereotaxic surgery system"? Is this for rodents or for primates or for humans? I think that stereotaxic surgery is difficult enough without being automated. Automation here would be a whole new set of problems. Besides, it would take the beauty out of learning this procedure. I am trying to imagine how this would work and I have difficulty imagining this due to the nature of the procedure.