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146 protocols using «model 1900»

1

Stereotaxic Viral Injection in Mice

2025
Mice were anaesthetized using isoflurane (Cp-pharma) and placed on a heating pad on a stereotaxic frame (Model 1900—Kopf Instruments). Carprofen (Rimadyl—Zoetis) (20 mg/kg body weight) was given via subcutaneous injection. Mice were bilaterally (or unilaterally for calcium imaging experiments) injected using glass pipettes (#708707, BLAUBRAND intraMARK) with 0.3 µl of virus in the CeA by using the following coordinates calculated with respect to bregma: for the CeA and CeL: −1.20 mm anteroposterior, ±2.87 mm lateral, −4.65 to −4.72 mm ventral; for the CeM −1.155 mm anteroposterior, ±2.87 mm lateral, −4.65 to −4.72 mm ventral. The following stereotaxic coordinates were used for the PBN: −5.2 mm anteroposterior, ±1.4 mm lateral, −3.85 mm ventral. Virus was allowed to be expressed for a minimum duration of 3 weeks before histology or behavioral paradigms. For animals not undergoing implant surgery, the incision was sutured.
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2

In Vivo Fiber Photometry of Neuromodulator Dynamics

2025
After acclimatizing to the holding facility for 7–9 days, the animals were anesthetized in an induction chamber (4% isoflurane, Piramal Healthcare, Maharashtra, India) and mounted on a stereotaxic frame under sterile conditions (Kopf Instruments, Model 1900) where they were maintained at 1–2% isoflurane for the duration of the surgery. For in vivo fiber photometry experiments, adult mice were injected bilaterally with 800 nl in each side of AAV5/Syn-FLEX-ChrimsonR-td-Tomato (Addgene) into the LC (AP: − 5.45 mm, ML: ± 1.25 mm, DV: − 3.8 mm) using a Hamilton syringe with a beveled needle and injected unilaterally with 400 nl of AAV1-hsyn-GRAB-NE2m or AAV9-hsyn-GRAB-DA2m (Yulong Li Lab) into the DG (AP: − 2.15 mm, ML: + 1.4 mm, DV: − 2.1 mm) using a Hamilton syringe with a blunted needle. Transgenic controls were injected bilaterally with 800 nl of AAV5/Syn-FLEX-ChrimsonR-td-Tomato into the LC (AP: − 5.45 mm, ML: ± 1.25 mm, DV: − 3.8 mm) using a Hamilton syringe with a beveled needle and injected unilaterally with 400 nl of AAV5-Ef1a-DIO-EYFP (Addgene) into the DG using a Hamilton syringe with a blunted needle. Mice then received intracranial fiber photometry implants in the DG (AP: − 2.15 mm, ML: + 1.4 mm, DV: − 1.7 mm), which were secured using MetaBond (C & B MetaBond). Mice were allowed to recover for at least six weeks following infusion of virus and intracranial implant prior to further behavioral testing or perfusion for projection mapping to ensure optimal viral expression. Mice were perfused at the conclusion of behavior to ensure optimal viral expression and optical implant placement location (Supplementary Fig. 6f).
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3

Stereotaxic Viral Delivery and Optogenetics in Mice

2024
4-5-week-old mice were deeply anesthetized using 3-4% vaporized Isoflurane and head-fixed on a stereotaxic apparatus (David Kopf Instruments Model 1900) for all stereotaxic surgeries. Anesthesia was maintained throughout the procedure at a level that prevented reflex response to a tail/toe pinch, using a continuous flow of 1-2% vaporized Isoflurane. Eye drops (Puralube Vet Ointment, Fisher Scientific, 2024927) were placed in each eye to prevent them from drying out, and vitals were checked every 10 min. An incision was made to expose the bregma and lambda point of the skull. A 1 mm drill was used to perforate the skull at the desired coordinates. Stereotaxic coordinates for the injection sites were determined using the Paxinos Brain atlas and adjusted experimentally. Using a syringe pump (PHD Ultra™, Harvard apparatus, no. 70-3007) installed with a microliter syringe (Hampton, 1482452A) and capillary glass pipettes with filament (Warner Instruments, G150TF-4), we infused the brain with 500 nl (for injections in the PL) or 1 μl (for injections in the VTA) of AAV solution for each injection site at a rate of 100 nl/min. To guarantee sufficient AAV expression across the anterior-posterior (AP) extent of the PL the AVV solutions were injected bilaterally at 2 injection sites for each brain hemisphere (from bregma: anterior–posterior (AP), +2.65 mm and +2.25 mm from bregma; mediolateral (ML), ±0.5 mm; dorsal–ventral (DV), −0.8 mm and -1.1 mm from the dura).
After injection of the PL, the pipette was left in place for 8 min to allow diffusion of the virus. When the surgery was completed, the scalp was disinfected with betadine and sutured with tissue adhesive glue (Vetbond tissue adhesive, 1469SB). For post-op pain treatment, mice received an injection of Buprenorphine SR (0.5 mg/kg) or Ethiqa XR (3.25 mg/kg).
When targeting the VTA, we injected the AAV solutions bilaterally (from bregma: AP, −3.2 mm; ML, ±0.5 mm; DV, −4.0 mm from the dura). After injection of the VTA, the pipette was left in place for 16 min to allow diffusion of the virus. Mice that later received optogenetic VTA stimulation were implanted during the same surgical procedure with a fiber optic cannula with Ceramic Ferrule (RWD Life Sciences, R-FOC-F200C-39NA). We used the same coordinates used for AAV injection with the following modifications: the fiber was implanted at a 10° angle, and the DV coordinate was reduced to −3.9. For selective stimulation of VTA dopaminergic neurons projecting to the PL, AAV were injected into the VTA and fiber optic cannulas were bilaterally implanted over the PL at a 10° angle (from bregma: AP, 2.5 mm; ML, ±0.9 mm; DV, −0.7 mm from the dura). To secure the implants to the skull, the skull was covered with a layer of OptiBond XTR Primer (OptiBond XTR Bottle Primer - 5 ml Bottle. Self-Etching) followed by OptiBond XTR Bottle Universal Adhesive (OptiBond XTR Bottle Universal Adhesive 5 ml Bottle. Self-Etching, Light-Cure). Finally, a thick layer (up to 0.5 cm thick) of Nano-optimized Flowable Composite (Tetric EvoFlow A2 Syringe - Nano-optimized Flowable Composite 1–2 Gram) was used to create a scaffold and secure the optic fiber to the skull. Polymerization of OptiBond XTR Primer, Adhesive, and Flowable Composite was achieved with dental LED light (Fencia Premium Silver LED Light, 5 W).
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4

Intracranial Viral Infusion Targeting VTA

2024
Procedures for intracranial viral infusion targeting the VTA were described previously (DeBaker et al., 2023 (link)). In brief, mice (>50 d) were placed in a stereotaxic frame (Model 1900, David Kopf Instruments; Tujunga, CA) under isoflurane anesthesia. Bilateral microinjectors consisting of a 33-gage stainless steel tube within a shorter 26-gage stainless steel tube were attached to polyethylene-20 tubing affixed to a 10 μL Hamilton syringe. Microinjectors were lowered through bilateral burr holes in the skull to the VTA (A/P: −2.75 mm, M/L: ±0.55 mm, D/V: −5.0 mm). AAV vectors (400 nL) were infused at a rate of 100 nL/min and microinjectors were left in place for 5 min to reduce backflow along the injection track upon removal. Behavioral experiments occurred 2–3 wk. after viral infusion.
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5

Stereotaxic Viral Injection and GRIN Lens Implantation in Mice

2024
Buprenorphine (0.1 mg/kg) was subcutaneously injected ~30 min before surgery for analgesia. Then, mice were anesthetized with isoflurane (1.5−2.0 % maintenance) through the oxygen-enriched air (95 %, 1–3 l/min, Oxymat III, Weinmann). Anesthesia level was monitored via breathing rates and foot and tail reflexes before and during surgery. Mice were placed in a stereotaxic apparatus (Model 1900, Kopf Instruments), and their body temperature was maintained through a heating pad (Rodent warmer, 53800 M, Stoelting). Their eyes were covered by an eye protective cream (Bepanthen Augen und Nasensalbe, Bayer). A mixture of Lidocaine (10 mg/kg) and Ropivacaine (3 mg/kg) was injected under the skin over the skull for local anesthesia. Stereotaxic viral injections were performed as previously described5 (link). Briefly, a small craniotomy was performed above the medial geniculate body (MGB, AP: −3.28, ML: −1.9, DV: −3.0 mm) by using a stereotaxic drill (Model 1911, Kopf) with a burr drill bit (105-0135-225, Kyocera). A pulled glass pipette (2-000-001, Drummond Scientific) filled with AAV vector was slowly lowered into the brain with the help of a micropositioner (Model 2650, Kopf). AAV2/1.syn.jGCaMP7s56 (link) (Addgene, 104487-AAV1, ca 500 nl, diluted by sterile PBS, 1-2x) was injected into MGB with a pressure ejection system (Picospritzer, Parker). One to two weeks after viral injection, a gradient refractive index (GRIN) lens (0.6 mm diameter, 7.3 mm length or 1.0 mm diameter, 4 mm length, Inscopix) was implanted during the second surgery (anesthesia and analgesia, see above). 0.6 mm lenses were implanted as previously described5 (link). Briefly, a 0.8 mm diameter craniotomy was performed above MGB (drill: 105-0709.400, Kyocera) and a small track was cut with a 0.7 mm sterile needle. Next, the GRIN lens was slowly advanced into the brain using the Micropositioner (Model 2650). For the implantation of 1.0 mm diameter lenses, a 1.2−1.3 mm craniotomy was performed above MGB using a hand drill (503599, World Precision Instrument) with a burr drill bit (200 µm diameter, C1.104.002, Bösch Dental). Tissue above MGB was slowly aspirated through a sterile blunt needle (27 G, Endo irrigation cannula) connected to a suction system. Sterilized phosphate buffer saline (PBS) was used to irrigate the brain until the bleeding stopped around the aspirated site. Next, the 1.0 mm lens was slowly advanced into the brain with the micropositioner. Both, 0.6 and 1.0 mm lenses were fixed to the skull with light curable glue (Loctite 4305, Henkel). A custom-made head bar was attached to the skull next to the GRIN lens, and the skull was sealed with Scotchbond (3 M), Vetbond (3 M) and dental acrylic (Paladur, Kulzer, Orth Jet, Lang Dental and/or C&B Super-Bond, Sun Medical). Meloxicam (5 mg/kg) was injected subcutaneously after the surgery for post-operative analgesia.
For optogenetic experiments, either AAV2/5-hsyn-Jaws-KGC-GFP-ER257 (link) (Addgene, 65014-AAV5, ca 500 nl, diluted by sterile saline, 2x) or AAV2/5-hSyn-EGFP (Addgene, 50465-AAV5, ca 500 nl, diluted by sterile saline, 2x) was bilaterally injected into MGB (AP: −3.2, ML: ±2, DV: −3.0/−3.3 mm) using the same methods as described above. Following the AAV injection, optical fibers were bilaterally implanted above MGB (AP: −3.2, ML: ±2, DV: −3 mm) using the Micropositioner on the same surgery. Other surgical procedures including anesthesia and local analgesia were the same as for GRIN lens implantations. Systemic analgesia was provided via carprofen in the drinking water (0.067 mg/ml) from ca. 12–24 h pre-surgery to ca. 72 h post-surgery.
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Top 5 protocols citing «model 1900»

1

Optogenetic Manipulation of Dynorphin Neurons

Adult (25–35 g) male preprodynorphin-IRES-cre (dyn-Cre) mice were group-housed, given access to food pellets and water ad libitum, and maintained on a 12 hr:12 hr light:dark cycle (lights on at 7:00 AM). All animals were kept in a sound-attenuated, isolated holding facility in the lab 1 week prior to surgery, post-surgery, and throughout the duration of the behavioral assays to minimize stress. All procedures were approved by the Animal Care and Use Committee of Washington University and conformed to US National Institutes of Health guidelines. For surgery, mice were anesthetized in an induction chamber (4% Isoflurane) and placed into a stereotaxic frame (Kopf Instruments, Model 1900) where they were maintained at 1%–2% isoflurane. We performed a craniotomy and unilaterally injected, using a blunt needle (86200, Hamilton Company), 300 nl of AAV5-DIO-ChR2-eYFP or AAV5-DIO-eYFP (Hope Center Viral Vector Core, viral titer 2 × 1013 vg/ml) into either the dNAcSh (stereotaxic coordinates from bregma: +1.30 anterior-posterior [AP], ±0.5 medial-lateral [ML], −4.25 mm dorsal-ventral [DV]) or vNAcSh (stereotaxic coordinates from bregma: +1.30 [AP], ±0.5 [ML], −4.75 mm [DV]), followed by fiber optic implantation (Sparta et al., 2012 (link)). We secured the implants using two bone screws and a dental dental cement headcap (Lang Dental). Mice were allowed to recover for 3 weeks prior to behavioral testing, permitting optimal expression of ChR2 in the Dyn-Cre cell bodies. For NorBNI local Infusion, mice were locally injected with NorBNI in the vNAcSh (2.5 μg/1 μl) and implanted with a fiber optic 2 weeks prior to behavior. These mice were allowed at least 1 week to recover post-surgery before beginning experimentation, well within the limits of norBNI antagonism.
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2

Surgical Procedures for Head-Fixed Mouse Electrophysiology

Recording from head-fixed mice involves two survival surgical procedures: first, to attach head fixation bars and optionally inject virus for optogenetics or fiber photometry; second, to prepare a craniotomy for inserting the microprobe. Surgeries were performed under a stereo microscope (Seiler iQ). All procedures were approved by the University of California, Los Angeles Chancellor’s Animal Research Committee. Male C57BL/6J, 8 – 12 week old mice were obtained from The Jackson Laboratory (stock no. 000664). Animals were maintained on a 12 hr light cycle, and group housed until the first surgery. Animals underwent the first aseptic surgery under isoflurane anesthesia on a stereotaxic instrument (Model 1900, Kopf Instruments). A stainless steel head fixation bar (online file repository, laser cut at Mainstay Manufacturing), was attached to each side of the skull with dental cement (Metabond, Parkell). Using two separate pieces helps reduce weight and improves access to the top of the skull for microprobe insertion (figure 7A). For experiments involving optogenetics or photometry, adeno-associated virus (AAV) was obtained from the University of North Carolina Vector Core. Craniotomies were made with a surgical drill, 0.3 mm drill bit diameter (X000QVJVF9, Daewon Industries). For optogenetics, AAV for the either the excitatory opsin AAV5/CaMKIIa-hChR2-eYFP or inhibitory opsin AAV5/CaMKIIa-eNpHR3.0-eYFP was injected in the secondary motor cortex (M2, coordinates relative to bregma: 2.5 mm anterior, 1.5 mm lateral, 1.3 mm ventral) at a volume of 300 nL by pulled glass pipettes (Nanoject II, Drummond Scientific). For photometry, 300 nL of AAVDJ/hsyn-GCaMP6s was injected in the same coordinates. The exposed skull was protected by covering with silicone elastomer (Kwik-Cast, World Precision Instruments) and then dental cement (Metabond, Parkell). A daily carprofen injection (5 mg/kg, s.c.) was administered for the first three days post-operatively, and analgesics (ibuprofen) and antibiotics (amoxicillin) were administered in the drinking water for the first week post-operatively. All mice were individually housed after surgery, and were allowed to recover for at least 2 weeks (3 weeks for animals injected with virus). The second surgery was also performed on stereotaxic equipment under isoflurane anesthesia. This procedure involved using a surgical drill to remove the protective dental cement, then tweezers to remove the Kwik-Cast layer to expose the skull, making a rectangular craniotomy above the recording site, and gently removing the dura to facilitate device insertion. An additional craniotomy was made over the posterior cerebellum to accommodate an electrical reference wire. The craniotomies were temporarily sealed with Kwik-Cast for the duration of the recovery period. Mice recovered for 6 – 10 hrs in their cage with unrestricted water access before being attached to a head bar holder (online file repository, manufactured at the UCLA Physics Machine Shop), and transferred to the recording setup.
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3

Targeted Viral Transduction and Optogenetic Manipulation

After the mice were acclimated to the holding facility for at least seven days, the mice were anaesthetized in an induction chamber (1-4% isoflurane) and placed into a stereotaxic frame (Kopf Instruments, Model 1900) where they were maintained at 1-2% isoflurane. Mice were then injected unilaterally or bilaterally, depending on experimental paradigm, using a blunt needle (86200, Hamilton Company) at a rate of 100 nL / min. The type of virus, injection volume, and stereotaxic coordinates for each experiment are listed in Supplementary Table 1. Mice were allowed to recover for five weeks prior to behavioral testing, permitting optimal expression of the virus. For optogenetic experiments, one week prior to behavioral testing, intracranial optic fiber implants were directed above the VTA (AP −3.15, ML ± 0.5, DV −4.25). Implants were secured using at least two bone screws and a dental cement headcap (Lang Dental).
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4

PKCε Knockdown in Amygdala Modulates Alcohol Consumption

To examine the role of PKCε in alcohol consumption, we used RNA interference to selectively reduce levels of PKCε in the amygdala. The design and validation of the lentiviruses used here has been described previously (Lesscher et al., 2008 (link)). Briefly, short hairpin RNA constructs were designed and incorporated into a pLentiLox 3.7 vector (kindly provided by L. van Parijs, MIT, Cambridge, MA) (Rubinson et al., 2003 (link)). Lentivirus was produced using Virapower packaging vectors (Invitrogen Carlsbad, CA, USA) for long-lasting PKCε knockdown in vivo. The shRNA construct used for this study was selected out of three constructs based on a high PKCε knockdown efficiency (>60%) in Neuro2A cells and in vivo in the amygdala (Lesscher et al., 2008 (link)). A sequence that did not recognize any known mammalian gene in a BLAST search was used as a control.
For microinjection of lentivirus, male F2 C57BL/6J ×129S4/SvJae wild type mice were anaesthetized with xylazine (7 mg/kg i.p.) and ketamine (100 mg/kg i.p.) and placed in a digital stereotaxic alignment system (Model 1900, David Kopf Instruments, Tujunga, CA, USA). The injectors (33 gauge, 0.2 mm outside diameter) were targeted to the central nucleus of the amygdala using the coordinates: −0.85 mm posterior to bregma, +/− 3.1 mm lateral to midline and −4.9 mm ventral from bregma. Lentivirus (2 μl, 80 × 106 pg p24 antigen/ml) was infused at a rate of 0.2 μl/min. The mice were allowed to recover for two weeks post-surgery. The animals used in this study had been used in a previous study (Lesscher et al 2008 (link)) examining the role of amygdala PKCε in anxiety-like behavior. No pharmacological treatments were given in the previous study and the animals were ethanol-naïve at the beginning of the current study. At the completion of the anxiety tests in the previous study, mice were adapted to the altered light-dark cycle (12:00 PM lights off) for two weeks.
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5

Stereotaxic Surgery for Implanting Guide Cannulae

Guide cannulae, stylets and tubing to make injection cannulae were obtained from Small Parts Inc. (Miami Lakes, FL). Guide cannulae were made of 25-gauge stainless steel tubing, pre-cut to 15.5mm. Stainless steel wire (0.0095 inch) was used to make stylets, which were inserted inside the guide cannulae to prevent obstruction. The guide cannulae were implanted 3mm above the VTA by stereotaxic surgery (Model 1900; David Kopf Instruments, Tujunga, CA) as previously described [33 (link), 34 (link)]. Mice were first anesthetized using a ketamine/xylazine cocktail (containing 100mg of Ketamine and 10mg of xylazine in 10mL saline), administered intraperitoneally in a volume of 0.1mL/10g of mouse weight. The animal’s dorsal scalp was shaved and a midline incision was made, revealing the skull from bregma to lambda (about 3 mm wide). The skull was cleaned with surgical scrub and sterilized using 100% ethanol (Pharmco Inc, Brookfield, CT). The 1997 edition of Franklin and Paxinos’ Mouse Brain Atlas was used for coordinates. The anterior-VTA coordinates (from bregma: caudal 3.16mm, lateral 0.5mm, and ventral 2.0mm) and posterior-VTA coordinates (from bregma: caudal 3.64mm, lateral 0.5mm, and ventral 2.0mm) were adjusted for each mouse. To accomplish this, the published distance between bregma and lambda in this species (4.21mm) was divided by the distance between bregma and lambda measured for each individual mouse. This quotient was then multiplied to each anterior or posterior coordinate listed above. Bilateral craniotomy holes were drilled at these individualized coordinates for placement of the guide cannulae. A third hole was drilled for an anchor screw (this third hole was enlarged using an 1/8 inch hand-held drill; Small Parts, Miami Lakes, FL, USA). Guide cannulae were lowered into position (2mm above VTA region of interest) and durelon carboxylate cement (Norristown, PA, USA) was applied to the exposed cranium to hold the assembly in place. Animals remained in the stereotaxic apparatus until cement dried. Following surgery, mice were subcutaneously administered the anti-inflammatory drug carprofen (Rimadyl, 5mg/kg, Pfizer Animal Health, USA), and the analgesic drug buprenorphine (Buprenex, 0.06mg/kg, Reckitt and Coleman Pharmaceuticals, Richmond, VA) and were monitored for healthy recovery.
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