Tamara Zeric for helping to obtain the multiphoton image in this paper. Supported by the Brain &#38; Behavior, Parkinson&#39;s and JPB Foundations, R01 MH108186 and R01 DA07418. F31 fellowship 1F31MH109293-01A1 to S C.

All procedures involving animals were performed according to the standards set by the Columbia University Medical Center Institutional Animal Care and Use Committee (IACUC). Euthanasia was performed via cervical dislocation under anesthesia with ketamine 100 mg/kg and xylazine 10 mg/kg injected intraperitoneally. All surgical procedures were performed in a sterile manner (surgeon was wearing head cap, face mask, sterile gloves, and clean disposable lab coat) and surgical tools were autoclaved between each use.
1. Presurgical Steps
<ol>
	<li>Autoclave all surgical instruments to ensure sterility.</li>
	<li>Anesthetize C57bl6 mice with isoflurane (4% for induction and 1.5-2% for surgery). Pinch the hind toe every 10 min to ensure proper depth of anesthesia. Increase anesthesia if vocalization or hind limb retraction is seen.</li>
	<li>Apply veterinary eye lubricant to the eyes to prevent drying.</li>
	<li>Remove the hair over the scalp with a 3 min application of hair removal gel or by shaving with an electric razor.</li>
	<li>Place the mouse in the stereotaxic frame on a heating pad or calibrated water recirculating blanket set to 38&#176;C.</li>
	<li>Administer buprenorphine (0.1 mg/kg) subcutaneously to provide presurgical analgesia.</li>
	<li>Disinfect the scalp with three alternating applications of chlorhexidine or betadine and 70% ethanol using a cotton tipped applicator.</li>
	<li>Inject 100 mL of bupivacaine diluted 50% in saline under the scalp to provide local anesthesia.</li>
</ol>
2. Piezoelectric Cranial Window Surgery
<ol>
	<li>Using autoclaved sterile surgical instruments, remove a 1 cm circle of scalp over the skull by cutting with surgical scissors in a circular pattern. Lift the scalp off the skull exposing the periosteum tissue over the bone.</li>
	<li>Use the tip of forceps to scrape the exposed bone clear of any remaining periosteum tissue. This is important to ensure that the dental cement will not fall off later.</li>
	<li>Set the piezoelectric surgery unit to vibrate on the lowest setting. Note that higher settings may break blood vessels due to intense vibrations.</li>
	<li>Fix the sterile circular tip to the handpiece. 
	NOTE: The 4 mm circular tip is well suited for surgeries. The large tip size contributes to the speed of the surgery as the entire window can be thinned simultaneously.</li>
	<li>Fill a 10 mL syringe with ice cold artificial cerebrospinal fluid (ACSF) and hold it in the hand not holding the piezoelectric handpiece.</li>
	<li>Irrigate the skull by dripping the ACSF from the syringe onto the skull at a rate of 1 mL/min. Alternatively, use a peristaltic pump positioned over the skull to free the use of the other hand. 
	NOTE: Irrigation with cold ACSF is important to prevent overheating due to the friction from the high frequency vibrations.</li>
	<li>Gently apply the vibrating surgical tip to the skull with a light circular motion (3 circles per 1 s) and thin the bone to the desired depth to perform a thin skull preparation (5 to 10 min). 
	NOTE: Here, 4 mm windows were made that were thinned to 20 &#181;m. However, the size of the window will vary with the size of the surgical tip. The thickness of the bone can be adjusted to the desired depth based on the length of time spent applying the tip to the skull.</li>
	<li>Gently adjust the angle of the handpiece and change the angle of the tip while applying pressure to the skull to thin the skull uniformly.</li>
	<li>To perform a complete bone removal without leaving any thin bone behind, thin the bone until cracks are visible around the window area. Remove the remaining flakes of thin bone with forceps without damaging the underlying dura.</li>
	<li>Soak a 1 mm piece of hemostatic collagen foam in cold ACSF. Hold the wet foam with forceps and place it on the thin skull. Allow it to sit on the thin skull&#160;for about 30 s to stop any micro bleeds from developing.</li>
	<li>Use forceps to place a 4 mm glass coverslip laterally over the window that has been thinned into the bone.</li>
	<li>Use the back of a small plastic or wooden cotton tipped applicator to apply approximately 100 &#181;L of dental acrylic to the skull to hold the window in place.</li>
</ol>
3. Post Surgical Care
<ol>
	<li>Remove the mouse from the surgical apparatus and place it in a heated cage for recovery. Monitor it continuously until it has regained full consciousness (assessed by normal mobility, gait, and behavior).</li>
	<li>Assess the animals twice a day for three days for manifestations of post-surgical pain including decreased activity, grooming, food, and water consumption, guarding behavior (e.g. limping or a hunched posture) or increased aggression. Do not return the animal to the company of other animals until it has fully recovered.</li>
	<li>Administer Buprenorphine (0.05&#8211;0.1 mg/kg) subcutaneously as an analgesic 8&#8211;12 h after the surgery for 3 days. Monitor the area of the window closely following surgery for dehiscence, inflammation, and signs of infection. If the animal does not return to baseline feeding and grooming behaviors, consult a veterinarian regarding possible interventions or euthanasia.</li>
</ol>

<ol>
	<li>Holtmaat, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term,+high-resolution+imaging+in+the+mouse+neocortex+through+a+chronic+cranial+window.">Long-term, high-resolution imaging in the mouse neocortex through a chronic cranial window.</a> Nat Protoc. 4, 1128-1144 (2009).</li><li>Yang, G., Pan, F., Parkhurst, C. N., Grutzendler, J., Gan, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thinned-skull+cranial+window+technique+for+long+term+imaging+of+the+cortex+in+live+mice.">Thinned-skull cranial window technique for long term imaging of the cortex in live mice.</a> Nat protoc. 12, 213-220 (2010).</li><li>Dorand, D. R., Barkauskas, D. S., Evans, T. A., Petrosiute, A., Huang, A. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+intravital+thinned+skull+and+cranial+window+approaches+to+study+CNS+immunobiology+in+the+mouse+cortex.">Comparison of intravital thinned skull and cranial window approaches to study CNS immunobiology in the mouse cortex.</a> Intravital. 3, e29728-1-e29728-8 (2014).</li><li>Drew, P. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chronic+optical+access+through+a+polished+and+reinforced+thinned+skull.">Chronic optical access through a polished and reinforced thinned skull.</a> Nat Methods. 7, 981-984 (2010).</li><li>Wang, J., Zhang, Y., Xu, T. H., Luo, Q. M., Zhu, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+innovative+transparent+cranial+window+based+on+skull+optical+clearing.">An innovative transparent cranial window based on skull optical clearing.</a> Laser Physics Letters. 9, (2012).</li><li>Yang, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Skull+Optical+Clearing+Solution+for+Enhancing+Ultrasonic+and+Photoacoustic+Imaging.">Skull Optical Clearing Solution for Enhancing Ultrasonic and Photoacoustic Imaging.</a> IEEE Trans Med Imaging. 35, (2016).</li><li>Vercellotti, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Piezoelectric+surgery+in+implantology:+a+case+report--a+new+piezoelectric+ridge+expansion+technique.">Piezoelectric surgery in implantology: a case report--a new piezoelectric ridge expansion technique.</a> Int J Periodontics Restorative Dent. 4, 358-365 (2000).</li><li>Vercellotti, T., De Paoli, S., Nevins, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+piezoelectric+bony+window+osteotomy+and+sinus+membrane+elevation:+introduction+of+a+new+technique+for+simplification+of+the+sinus+augmentation+procedure.">The piezoelectric bony window osteotomy and sinus membrane elevation: introduction of a new technique for simplification of the sinus augmentation procedure.</a> Int J Periodontics Restorative Dent. 6, 561-567 (2001).</li><li>Vercellotti, T., Podesta, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Orthodontic+microsurgery:+a+new+surgically+guided+technique+for+dental+movement.">Orthodontic microsurgery: a new surgically guided technique for dental movement.</a> Int J Periodontics Restorative Dent. 4, 325-331 (2007).</li><li>Stubinger, S., Stricker, A., Berg, B. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Piezoelectric+surgery+in+implant+dentistry.">Piezoelectric surgery in implant dentistry.</a> Clin Cosmet Investig Dent. 7, 115-124 (2015).</li><li>Basheer, S. A., Govind, R. J., Daniel, A., Sam, G., Adarsh, V. J., Rao, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparative+Study+of+Piezoelectric+and+Rotary+Osteotomy+Technique+for+Third+Molar+Impaction.">Comparative Study of Piezoelectric and Rotary Osteotomy Technique for Third Molar Impaction.</a> J Contemp Dent Pract. 18, 60-64 (2017).</li><li>Pavlikova, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Piezoelectric+surgery+prevents+brain+tissue+damage:+an+experimental+study+on+a+new+rat+model.">Piezoelectric surgery prevents brain tissue damage: an experimental study on a new rat model.</a> Int J Oral Maxillofac Surg. 40, 840-844 (2011).</li></ol>

There are no disclosures.

The surgery can be modified by changing the surgical tip. There are many different sizes of tip that can be applied to the handpiece. Changing the size or shape of the tip will result in different sized windows. In addition to the 4 mm tip, we also tried a 3 mm tip and found it also worked well. Even with irrigation with ice cold ACSF, we were not able to get good results with tips that were too narrow (approximately 0.25 mm) due to the concentrated vibrations causing too much heat and burning the bone leading to damage to the underlying dura. We have not tried the multitude of other tips that are available, and we anticipate that other labs may find new applications for these different tips. While the surgery is relatively straightforward, we found that there are several steps that may require troubleshooting. The first is that the vibrating tip produces a lot of turbulence in the ACSF, which diminishes visibility and produces difficulty in ascertaining bone depth during the bone thinning step of the surgery. We recommend that if it is too difficult to see how thin the bone is, take a break to dry the skull and window by applying a sterile cotton swab to the side of the bone. Do not apply the cotton swab directly to the window, since the rough surface is damaging to thin bone. After checking the depth, reapply ACSF and continue with the surgery. We also found that if there is damage to the dura, it is usually due to the surgeon putting too much pressure on the tip. Holding the handpiece more gently and applying less force will likely fix this problem. If the surgeon is forcibly scraping the tip on the bone, it will cause a break in the thin bone and damage the skull. Finally, if after thinning the bone, the underlying dura appears bruised, it is likely due to excess heat from the vibrations. Increasing the rate of ACSF flow will fix this issue.
The main limitation of PHASOR is that the handpiece can thin the bone, but cannot remove all the bone over the dura. The diamond coated tip has small rough bumps. The rubbing needed to thin the skull would abrade the dura and cause bleeding if it were used to remove all the bone. Thus, the use of forceps is required to remove the remaining thin layer. While there is ongoing debate on whether imaging through a thin skull produces less inflammation and less proliferation of microglia and reactive astrocytes, in some cases a window with no bone remaining is preferred, e.g., to provide a greater depth of imaging3.
We have optimized cranial window preparation for multiphoton imaging in mice. PHASOR is, to our knowledge, the first application of piezoelectric technique to rodent cranial window surgery for optical imaging. We find that the use of piezoelectric surgery in mice shares the advantages of increased speed and decreased adverse events also reported in humans7,8,9,10. Utilization of the piezoelectric device resulted in faster surgeries (Figure 2A) and fewer adverse events of bleeding when compared to a high-speed rotary drill (Figure 2B). We also found, within our lab, that PHASOR was easier for new surgeons to learn than traditional approaches to cranial window surgery. Advantages of speed and ease of use are likely to differ among surgeons. Thin skull preparations typically require 30-45 min to thin the bone with a drill and scalpel2, while the PHASOR approach typically requires less than 10 min.
We found that in windows prepared with PHASOR we could image calcium transients in layer 4 pyramidal neurons in motor cortex that had been transfected with adeno associated virus encoding the calcium indicator JRGECO1a (AAV9.Syn.NES-jRGECO1a.WPRE.SV40). We also found that, compared to a thin skull cranial window prepared with a rotary drill and scalpel, the imaging region was larger, while a thin skull prepared with a rotary drill and scalpel has an imaging window of 20 &#181;m diameter2. With PHASOR, we were able to quickly thin an area of 3-4 mm. This is similar to the imaging window reported with PoRTS4. This larger window retains the benefit of rapid imaging reported for thin skull windows prepared with a dental drill and scalpel. Moreover, the window prepared with PHASOR can be immediately imaged, as opposed to traditional bone removal windows that may require weeks to heal before the window can be optimally imaged3.
This technique can be applied to study alterations in blood flow in the vessels under the dura. An important future study is to compare immunoreactivity of PHASOR with other methods of cranial window. This would be important to determine if PHASOR produces less inflammation in comparison to existing methods. We hope that the piezoelectric surgical technique documented here will allow more labs to perform cranial windows and successfully use multiphoton imaging in vivo.
Make sure that the handpiece is set to vibrate on the lowest setting and that the ACSF is being irrigated at a constant rate. The ice cold ACSF must be applied constantly or it will heat up and damage the dura. We found that ACSF must be applied at a rate of at least 1 mL/min. Either use a syringe held in the hand, or use a peristaltic pump, rather than the built-in irrigation in the handpiece. The irrigation system in the handpiece ejects ACSF too forcefully and produces too much turbulence, which will significantly impair visibility.

Cranial window surgery to prepare rodents for multiphoton imaging in vivo has become an important technique in neuroscience. The removal or thinning of bone is necessary to prepare the mouse for optical imaging with a multiphoton microscope. This surgery is performed either by completely removing an area of bone to expose the underlying dura1, or by thinning a region of bone without full removal from the dura2. The thin skull approach may produce less inflammation and activation of microglia3 but provides a shallower depth of imaging, a smaller imaging window size (200 &#181;m) and a limited time period during which the window can be imaged due to bone regrowth2. The addition of a polished and reinforced glass window (PoRTS) can increase the imaging size and imaging period but is difficult to perform4.
Both current surgeries use a high speed rotary drill to thin or remove the bone from the skull. The thin skull technique also uses a scalpel after the drill to further thin the bone2. The PoRTS technique requires the extra step of high speed polishing with grit4. In a high speed rotary drill, an air powered turbine or electric motor causes the drill bit to spin at a high velocity. As rotary drills section both bone and soft tissue, there is a risk of damaging the dura and underlying blood vessels. The success of the surgery depends on the skill of the surgeon. In addition to these windows prepared with mechanical surgical methods, a chemical method of optically clearing the skull with different solutions has been developed5,6. However, since piezoelectric surgery is a mechanical method of surgery, our comparisons here will be limited to other mechanical methods.
Piezoelectric surgical devices utilize ultrasonic vibrations to break down mineralized bone without damaging underlying soft tissue, and thus offer an approach to rapidly thin a large area of bone. In a piezoelectric surgical handpiece, the turbine is replaced by a stack of ceramic disks and when current is applied, the disks vibrate at ultrasonic frequencies. The vibrations are transferred through the handpiece to diamond coated tips to cut through bone without damaging soft tissues, an advantage over rotary drills that do not discriminate between tissue types. Piezoelectric surgery was originally developed for use in human by Tomaso Vercellotti and has led to improvements in dental and cranio-maxilofacial surgery7,8,9,10,11.
Piezoelectric surgery has been used to create an osteotomy in Wistar rats and was found by magnetic resonance imaging (MRI) and histology to produce significantly less damage than a traditional dental drill12. The authors concluded that piezoelectric surgery was safe for removing the bone near soft brain tissue. Mice, however, have a thinner dura that is more easily damaged, and that study did not prepare windows for chronic optical imaging. Chronic imaging requires that blood vessels are not damaged and that blood clots do not form under the window. Damage to the dura leads to inflammation that causes the window to cloud, and activates microglia and the proliferation of reactive astrocytes. Here we have optimized piezoelectric surgery for mice to create both thin skull and full bone removal cranial windows suitable for chronic imaging. We compared this surgical technique to cranial window surgeries prepared with a high speed rotary drill.

<table><tbody><tr><td>Piezosurgery Touch</td><td>Mectron</td><td>5120062</td><td>Piezosurgery GP model has the same settings</td></tr><tr><td>Circular 4mm flat piezosurgery tip (# OT11)</td><td>Mectron</td><td>3370019</td><td>This tip was ideal for our windows but there are many other tips of different sizes availible.</td></tr><tr><td>Stereotax frame</td><td>Kopf</td><td>963</td><td></td></tr><tr><td>Mouse adaptor</td><td>Stoelting</td><td>51625</td><td></td></tr><tr><td>Peristaltic pump for irrigation.</td><td>Cole-Parmer</td><td>WU-77120-42</td><td>Makes it easier to irrigate and frees up the other hand to provide stability. Irrigation can be performed by hand with a syringe if necessary.</td></tr><tr><td>Avitene Ultrafoam</td><td>Bard-Davol</td><td>1050020</td><td>Important to stop any minor bleeding instantly.</td></tr><tr><td>C&amp;amp;B Metabond</td><td>Parkell</td><td>S380</td><td>Much stronger than regular dental acrylic.</td></tr><tr><td>Artificial cerebro spinal fluid (ACSF)</td><td>Tocris</td><td>3525</td><td></td></tr><tr><td>Puralube opthalmic ointment</td><td>Dechra</td><td>17033-211-38</td><td></td></tr><tr><td>Mice</td><td>JAX</td><td>664</td><td></td></tr><tr><td>Prairie Ultima multiphoton microscope</td><td>Bruker</td><td></td><td></td></tr><tr><td>JRGECO1a (AAV9.Syn.NES-jRGECO1a.WPRE.SV40)</td><td>UPENN Vector Core</td><td></td><td></td></tr></tbody></table>

piezo high accuracy surgical osteal removal (phasor): a technique for improved cranial window surgery in mice

Multiphoton microscopy has been widely adapted for imaging neurons in vivo. Repeated imaging requires implantation of a cranial window or repeated thinning of the skull. Cranial window surgery is typically performed with a high speed rotary drill, and many investigators find it challenging to prevent the drill from damaging the delicate dura and blood vessels. Extensive training and practice is required to remove the bone without damage to underlying tissue and thus cranial window surgery can be difficult, time consuming, and produce tissue damage. Piezoelectric surgery, which is extensively used for maxillofacial and dental surgery, utilizes ultrasonic vibrations to remove bone without damaging soft tissues. We have developed a method applying piezoelectric surgery to improve cranial window surgery in mice in preparation for multiphoton imaging. Comparisons within our lab find that the method requires less surgery time and has a lower average rate of complications due to dural bleeding than cranial window surgery with a rotary drill.

Before proceeding with piezoelectric surgery, remove any residual periosteum from the skull. Once the skull is opaque and smooth (Figure 1a), the surgeon may begin piezoelectric surgery. When removing bone with the vibrating piezo tip, it is critical to irrigate the skull with ice cold ACSF. Proper irrigation is achieved when the bottom 1 mm of the tip is submerged in ACSF (Figure 1b). Without proper irrigation, the bone will overheat and damage the brain. After the skull has been thinned to the desired depth chosen by the surgeon, there may be some residual bleeding from blood vessels located in the bone. In order to rapidly cease all bleeding, apply a small 1 mm piece of collagen foam soaked in ACSF to the area of the window. Let the collagen sit on the skull for approximately 30 s (Figure 1c). After the collagen foam is removed, the window will appear translucent, allowing clear visualization of the blood vessels in the dura. The dura will be intact without significant bruising. (Figure 1d). If the dura appears red and inflamed, it is likely due to broken blood vessels from too much pressure applied to the tip during the surgery. To protect the new window and prepare it for chronic imaging, a glass coverslip must be placed over the area. A properly applied glass coverslip will gently sit on top of the window and will not cause any damage to the area. The glass coverslip must cover the entire window. If the glass coverslip is properly applied to the skull, the window will remain translucent under the glass. All the blood vessels in the dura will still be visible (Figure 1e). Dental acrylic must be applied around the glass coverslip to permanently adhere it to the surface of the skull. The edges of the glass coverslip must also be covered in dental acrylic (Figure 1f).
We found that piezoelectric surgery is typically much faster than a traditional cranial window, taking around 10-12 min per surgery (Figure 2A). We also found that there were less complications due to dura bruising and bleeding as observed by eye (Figure 2B). A properly prepared window will allow multiphoton imaging of fluorescent indicators in cortical neurons in vivo. We chose to image the red calcium indicator JRGECO1a in cell bodies of layer 4 cortical neurons (Figure 3A-3B). We could observe calcium transients in these cell bodies through the window prepared using PHASOR.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56172/56172fig1.jpg" /> 
Figure 1: Piezoelectric surgery. (a) The skull after it has been properly prepared for PHASOR. All the periosteal tissue has been removed, leaving a smooth and clean surface. (b) A representative image of PHASOR in progress. The bottom 1 mm of the vibrating surgical tip is submerged in ice cold ACSF. The fluid is applied to the skull at a rate of 1 mL/min. A light circular motion is applied to the skull and the bone is thinned via ultrasonic vibrations. (c) After the skull has been thinned, a 1 mm circular piece of collagen foam soaked in cold ACSF is allowed to rest on the window to stop any micro bleeds from the bone. (d) A successful window with a translucent dura displays all of the intact blood vessels. There is no visible damage or bruising to the surface of the brain. (e) A glass coverslip is placed over the window to protect the surface of the brain. (f) Dental acrylic is applied to the skull to permanently fix the glass coverslip over the window. Scale bar 1 mm in all images.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56172/56172fig2.jpg" /> 
Figure 2. Comparison of success rate in training for PHASOR or dental drill surgery. (A) Average time per surgery was lower for PHASOR than dental drill surgeries (p&#60;0.05, two tailed t test) n= 30 surgeries per group and average 10 surgeries per surgeon. (B) The percentage of successful surgeries (defined as no bleeding or visible damage to the dura as observed by the surgeon&#39;s eye through the objective set to 350X zoom) was higher for PHASOR than dental drill surgeries (n = 30 surgeries for each group, one tailed z test). Error bars show SD.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56172/56172fig3.jpg" /> 
Figure 3. Multiphoton imaging in vivo through a cranial window thinned with PHASOR. (A) Calcium transients observed with the red calcium indicator JRGECO1a were imaged in layer 4 pyramidal neurons in murine motor cortex in vivo. Scale bar = 100 &#181;m. (B) The same window but different region of layer 4 motor cortex. Headfixed mouse with 3 mm window prepared with PHASOR. Scale bar = 30 &#181;m. Imaged with a multiphoton microscope, 40X objective, 1040 nm excitation wavelength.

Watch this Scientific Journal Video about Piezo High Accuracy Surgical Osteal Removal PHASOR: A Technique for Improved Cranial Window Surgery in Mice at JoVE.com

Piezo High Accuracy Surgical Osteal Removal PHASOR: A Technique for Improved Cranial Window Surgery in Mice

Piezoelectric surgery has led to improvements in human maxillofacial and dental surgery. We have developed a protocol to optimize piezoelectric surgery for cranial window surgery in mice.

Piezo High Accuracy Surgical Osteal Removal (PHASOR): A Technique for Improved Cranial Window Surgery in Mice

Multiphoton microscopy has been widely adapted for imaging neurons in vivo. Repeated imaging requires implantation of a cranial window or repeated ...

piezo-high-accuracy-surgical-osteal-removal-phasor-technique-for

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

10.7K Views.  Columbia University Medical Campus. Piezoelectric surgery has led to improvements in human maxillofacial and dental surgery. We have developed a protocol to optimize piezoelectric surgery for cranial window surgery in mice.

Video: Piezo High Accuracy Surgical Osteal Removal PHASOR: A Technique for Improved Cranial Window Surgery in Mice

Imaging Glioma Initiation In Vivo Through a Polished and Reinforced Thin-skull Cranial Window

Glioma is the one of the most lethal forms of human cancer. The most effective glioma therapy to date-surgery followed by radiation treatment-offers patients only modest benefits, as most patients do not survive more than five years following diagnosis due to glioma relapse 1,2. The discovery of cancer stem cells in human brain tumors holds promise for having an enormous impact on the development of novel therapeutic strategies for glioma 3. Cancer stem cells are defined by their ability both to self-renew and to differentiate, and are thought to be the only cells in a tumor that have the capacity to initiate new tumors 4. Glioma relapse following radiation therapy is thought to arise from resistance of glioma stem cells (GSCs) to therapy 5-10. In vivo, GSCs are shown to reside in a perivascular niche that is important for maintaining their stem cell-like characteristics 11-14. Central to the organization of the GSC niche are vascular endothelial cells 12. Existing evidence suggests that GSCs and their interaction with the vascular endothelial cells are important for tumor development, and identify GSCs and their interaction with endothelial cells as important therapeutic targets for glioma. The presence of GSCs is determined experimentally by their capability to initiate new tumors upon orthotopic transplantation 15. This is typically achieved by injecting a specific number of GBM cells isolated from human tumors into the brains of severely immuno-deficient mice, or of mouse GBM cells into the brains of congenic host mice. Assays for tumor growth are then performed following sufficient time to allow GSCs among the injected GBM cells to give rise to new tumors-typically several weeks or months. Hence, existing assays do not allow examination of the important pathological process of tumor initiation from single GSCs in vivo. Consequently, essential insights into the specific roles of GSCs and their interaction with the vascular endothelial cells in the early stages of tumor initiation are lacking. Such insights are critical for developing novel therapeutic strategies for glioma, and will have great implications for preventing glioma relapse in patients. Here we have adapted the PoRTS cranial window procedure 16and in vivo two-photon microscopy to allow visualization of tumor initiation from injected GBM cells in the brain of a live mouse. Our technique will pave the way for future efforts to elucidate the key signaling mechanisms between GSCs and vascular endothelial cells during glioma initiation.

Imaging Glioma Initiation In Vivo Through a Polished and Reinforced Thin-skull Cranial Window

By combining a polished and reinforced thin-skull (PoRTS) cranial window and glioblastoma (GBM) cell injection, we can observe glioma initiation and growth from injected GBM cells in the brain of a live mouse longitudinally.

Glioma is the one of the most lethal forms of human cancer. The most effective glioma therapy to date-surgery followed by radiation treatment-offers ...

Medicine

Thinned-skull Cortical Window Technique for In Vivo Optical Coherence Tomography Imaging

Optical coherence tomography (OCT) is a biomedical imaging technique with high spatial-temporal resolution. With its minimally invasive approach OCT has been used extensively in ophthalmology, dermatology, and gastroenterology1-3. Using a thinned-skull cortical window (TSCW), we employ spectral-domain OCT (SD-OCT) modality as a tool to image the cortex in vivo. Commonly, an opened-skull has been used for neuro-imaging as it provides more versatility, however, a TSCW approach is less invasive and is an effective mean for long term imaging in neuropathology studies. Here, we present a method of creating a TSCW in a mouse model for in vivo OCT imaging of the cerebral cortex.

Thinned-skull Cortical Window Technique for In Vivo Optical Coherence Tomography Imaging

We present a method of creating a thinned-skull cortical window (TSCW) in a mouse model for in vivo OCT imaging of the cerebral cortex.

Optical coherence tomography (OCT) is a biomedical imaging technique  with high spatial-temporal resolution. With its minimally invasive approach OCT  has ...

Real-Time Imaging of CCL5-Induced Migration of Periosteal Skeletal Stem Cells in Mice

Periosteal skeletal stem cells (P-SSCs) are essential for lifelong bone maintenance and repair, making them an ideal focus for the development of therapies to enhance fracture healing.&#160; Periosteal cells rapidly migrate to an injury to supply new chondrocytes and osteoblasts for fracture healing. Traditionally, the efficacy of a cytokine to induce cell migration has only been conducted in vitro by performing a transwell or scratch assay. With advancements in intravital microscopy using multiphoton excitation, it was recently discovered that 1) P-SSCs express the migratory gene CCR5 and 2) treatment with the CCR5 ligand known as CCL5 improves fracture healing and the migration of P-SSCs in response to CCL5. These results have been captured in real-time. Described here is a protocol to visualize P-SSC migration from the calvarial suture skeletal stem cell (SSC) niche towards an injury after treatment with CCL5. The protocol details the construction of a mouse restraint and imaging mount, surgical preparation of the mouse calvaria, induction of a calvaria defect, and acquisition of time-lapse imaging.

This protocol describes the detection of CCL5-mediated periosteal skeletal stem cell migration in real-time using live animal intravital microscopy.

Periosteal skeletal stem cells (P-SSCs) are essential for lifelong bone maintenance and repair, making them an ideal focus for the development of ...

Developmental Biology

The Mouse Round-window Approach for Ototoxic Agent Delivery: A Rapid and Reliable Technique for Inducing Cochlear Cell Degeneration

Investigators have utilized a wide array of animal models and investigative techniques to study the mammalian auditory system. Much of the basic research involving the cochlea and its associated neural pathways entails exposure of model cochleae to a variety of ototoxic agents. This allows investigators to study the effects of targeted damage to cochlear structures, and in some cases, the self-repair or regeneration of those structures. Various techniques exist for delivery of ototoxic agents to the cochlea. When selecting a particular technique, investigators must consider a number of factors, including the induction of inadvertent systemic toxicity, the amount of cochlear damage produced by the surgical procedure itself, the type of lesion desired, animal survivability, and reproducibility/reliability of results. Currently established techniques include parenteral injection, intra-peritoneal injection, trans-tympanic injection, endolymphatic sac injection, and cochleostomy with perilymphatic perfusion. Each of these methods has been successfully utilized and is well described in the literature; yet, each has various shortcomings. Here, we present a technique for topical application of ototoxic agents directly to the round window niche. This technique is non-invasive to inner ear structures, produces rapid onset of reliably targeted lesions, avoids systemic toxicity, and allows for an intra-animal control (the contra-lateral ear). Results stemming from this approach have helped deeper understanding of auditory pathophysiology, cochlear cell degeneration, and regenerative capacity in response to an acute injury. Future investigations may use this method to conduct interventional studies involving gene therapy and stem cell transplantation to combat hearing loss.

Various methods exist for introducing ototoxic agents to the cochleae of animal models. Presented is a surgical protocol for delivery of ototoxic agents to the round window niche. The procedure is reliable, creates targeted intra-cochlear lesions, and avoids mechanical damage to the microarchitecture. Examination of cochlear self-repair/regeneration is possible.

Investigators have utilized a wide array of animal models and investigative techniques to study the mammalian auditory system. Much of the basic research ...

Craniotomy Procedure for Visualizing Neuronal Activities in Hippocampus of Behaving Mice

Imaging neuronal activities at single-cell resolution in awake behaving animals is a very powerful approach for the investigation of neural circuit functions in systems neuroscience. However, high absorbance and scattering of light in mammalian tissue limit intravital imaging mostly to superficial brain regions, leaving deep-brain areas, such as the hippocampus, out of reach for optical microscopy. In this video, we show the preparation and implantation of the custom-made imaging window to enable chronic in vivo imaging of the dorsal hippocampal CA1 region in head-fixed behaving mice. The custom-made window is supplemented with an infusion cannula that allows targeted delivery of viral vectors and drugs to the imaging area. By combining this preparation with wide-field imaging, we performed a long-term recording of neuronal activity using a fluorescent calcium indicator from large subsets of neurons in behaving mice over several weeks. We also demonstrated the applicability of this preparation for voltage imaging with single-spike resolution. High-performance genetically encoded indicators of neuronal activity and scientific CMOS cameras allowed the recurrent visualization of subcellular morphological details of single neurons at high temporal resolution. We also discuss the advantages and potential limitations of the described method and its compatibility with other imaging techniques.

This article demonstrates the preparation of a custom-made imaging window supplemented with infusion cannula and its implantation onto the CA1 region of the hippocampus in mice.

Imaging neuronal activities at single-cell resolution in awake behaving animals is a very powerful approach for the investigation of neural circuit ...

Implantation of a Cranial Window for Repeated In Vivo Imaging in Awake Mice

To fully understand the cellular physiology of neurons and glia in behaving animals, it is necessary to visualize their morphology and record their activity in vivo in behaving mice. This paper describes a method for the implantation of a chronic cranial window to allow for the longitudinal imaging of brain cells in awake, head-restrained mice. In combination with genetic strategies and viral injections, it is possible to label specific cells and regions of interest with structural or physiological markers. This protocol demonstrates how to combine viral injections to label neurons in the vicinity of GCaMP6-expressing astrocytes in the cortex for simultaneous imaging of both cells through a cranial window. Multiphoton imaging of the same cells can be performed for days, weeks, or months in awake, behaving animals. This approach provides researchers with a method for viewing cellular dynamics in real time and can be applied to answer a number of questions in neuroscience.

Implantation of a Cranial Window for Repeated In Vivo Imaging in Awake Mice

Presented here is a protocol for the implantation of a chronic cranial window for the longitudinal imaging of brain cells in awake, head-restrained mice.

To fully understand the cellular physiology of neurons and glia in behaving animals, it is necessary to visualize their morphology and record their ...

Assessment of Thermal Damage from Robot-Drilled Craniotomy for Cranial Window Surgery in Mice

Cranial window surgery allows for the imaging of brain tissue in live mice with the use of multiphoton or other intravital imaging techniques. However, when performing any craniotomy by hand, there is often thermal damage to brain tissue, which is inherently variable surgery-to-surgery and may be dependent on individual surgeon technique. Implementing a surgical robot can standardize surgery and lead to a decrease in thermal damage associated with surgery. In this study, three methods of robotic drilling were tested to evaluate thermal damage: horizontal, point-by-point, and pulsed point-by-point. Horizontal drilling utilizes a continuous drilling schematic, while point-by-point drills several holes encompassing the cranial window. Pulsed point-by-point adds a &#34;2 s on, 2 s off&#34; drilling scheme to allow for cooling in between drilling. Fluorescent imaging of Evans Blue (EB) dye injected intravenously measures damage to brain tissue, while a thermocouple placed under the drilling site measures thermal damage. Thermocouple results indicate a significant decrease in temperature change in the pulsed point-by-point (6.90 &#176;C &#177; 1.35 &#176;C) group compared to the horizontal (16.66 &#176;C &#177; 2.08 &#176;C) and point-by-point (18.69 &#176;C &#177; 1.75 &#176;C) groups. Similarly, the pulsed point-by-point group also showed significantly less EB presence after cranial window drilling compared to the horizontal method, indicating less damage to blood vessels in the brain. Thus, a pulsed point-by-point drilling method appears to be the optimal scheme for reducing thermal damage. A robotic drill is a useful tool to help minimize training, variability, and reduce thermal damage. With the expanding use of multiphoton imaging across research labs, it is important to improve the rigor and reproducibility of results. The methods addressed here will help inform others of how to better use these surgical robots to further advance the field.

Cranial windows have become a ubiquitously implemented surgical technique to allow for intravital imaging in transgenic mice. This protocol describes the use of a surgical robot that performs semi-automated bone drilling of cranial windows and can help reduce surgeon-to-surgeon variability and partially mitigate thermal blood-brain barrier damage.

Cranial window surgery allows for the imaging of brain tissue in live mice with the use of multiphoton or other intravital imaging techniques. However, ...

Calvarial Window Generation in Mouse Model: A Surgical Procedure to Create Window in the Top Part of the Mouse Skull

Source: Geisler, J. A. et al. <a href="https://jove.unicartagenaproxy.elogim.com/v/61272/modeling-brain-metastases-through-intracranial-injection-magnetic">Modeling Brain Metastases Through Intracranial Injection and Magnetic Resonance Imaging.</a> J. Vis. Exp. (2020)
In this video, we describe the surgical procedure to create a calvarial window for accessing the brain of a mouse model.

Source: Geisler, J. A. et al. Modeling Brain Metastases Through Intracranial Injection and Magnetic Resonance Imaging. J. Vis. Exp. (2020)
In this video, ...

JoVE Encyclopedia of Experiments

Cancer Research

Cancers of the Nervous System

In vivo studies

Implantation of Cranial Imaging Window in Mouse Model: A Surgical Procedure to Implant Glass-based Coverslip for Stable Optical Access to Regions of Murine Brain

Source: Alieva, M. et al. <a href="https://jove.unicartagenaproxy.elogim.com/v/59278/longitudinal-intravital-imaging-brain-tumor-cell-behavior-response-to">Longitudinal Intravital Imaging of Brain Tumor Cell Behavior in Response to an Invasive Surgical Biopsy.</a>&#160;J. Vis. Exp.&#160; (2019)
This video presents a surgical procedure to implant cranial window in a mouse model for stable optical access to different regions of the brain.

Source: Alieva, M. et al. Longitudinal Intravital Imaging of Brain Tumor Cell Behavior in Response to an Invasive Surgical Biopsy.&#160;J. Vis. Exp.&#160; ...

Installation of Optical Windows in the Brain of Transgenic Mice to Image Neural Activities

Source: Lin, X., et al. <a href="https://appjove.unicartagenaproxy.elogim.com/v/56297">Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice.</a> J. Vis. Exp. (2019).
This video demonstrates the brain&#39;s neural activity recording through an optical window in transgenic mice. The mice express a fluorescent protein that enables the detection of neural activities. A small portion of the skull is carefully removed to install an optical window, and the skull is then covered with black cement to block light before imaging.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

Piezo High Accuracy Surgical Osteal Removal (PHASOR): A Technique for Improved Cranial Window Surgery in Mice

Summary

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Chapters in this video

Imaging Glioma Initiation In Vivo Through a Polished and Reinforced Thin-skull Cranial Window

Thinned-skull Cortical Window Technique for In Vivo Optical Coherence Tomography Imaging

Real-Time Imaging of CCL5-Induced Migration of Periosteal Skeletal Stem Cells in Mice

The Mouse Round-window Approach for Ototoxic Agent Delivery: A Rapid and Reliable Technique for Inducing Cochlear Cell Degeneration

Craniotomy Procedure for Visualizing Neuronal Activities in Hippocampus of Behaving Mice

Implantation of a Cranial Window for Repeated In Vivo Imaging in Awake Mice

Assessment of Thermal Damage from Robot-Drilled Craniotomy for Cranial Window Surgery in Mice

Calvarial Window Generation in Mouse Model: A Surgical Procedure to Create Window in the Top Part of the Mouse Skull

Implantation of Cranial Imaging Window in Mouse Model: A Surgical Procedure to Implant Glass-based Coverslip for Stable Optical Access to Regions of Murine Brain

Installation of Optical Windows in the Brain of Transgenic Mice to Image Neural Activities