Supported by DK065872 (ENP), a Smith Family Foundation Award of Excellence in Biomedical Research (ENP), F31 DA023760.

Summary
Two-month old average age C57BL/6J mice or equivalent or three-month old average age Sprague Dawley rats or equivalent are anesthetized with ketamine (60 mg/kg i.p. for rats; 100 mg/kg i.p for mice) and xylazine (10 mg/kg, i.p. for either species). Sedation is monitored using a gentle toe pinch withdraw reflex demonstrated in <a href="https://www.jove.com/index/details.stp?ID=236">Walantus et al.</a>(JoVE, 6, 2007) and <a href="https://www.jove.com/index/details.stp?ID=404">Szot et al.</a>(JoVE, 9, 2007).&nbsp; Thermoregulation can be provided through a thermostatregulated heating pad (ALA Instruments Inc.) and monitored through a rectal thermometer. Head is shaved of fur and cleaned with iodine before incision. After skin incision (2 cm long for rats, 1 cm long for mice) and removal of all soft tissue from the surface of the skull, placement of the guide cannula is determined in relation to bregma. A 6 mm hole is drilled through the skull with a battery-operated driller designed for rodent surgery (Fine Science Tools, Inc.). Care is taken so that the drill bit does not penetrate through meningeal membranes or blood vessels. Skull is implanted with bilateral 5 mm 21 gauge stainless steel guide shafts leading to the posterior nucleus accumbens, dorsal striatum or prefrontal cortex. The stereotaxic coordinates are established as per Franklin and Paxinos, 1997 (The Mouse Brain in Stereotaxic Coordinates, Academic Press) or Paxinos and Watson, 2006 (The Rat Brain in Stereotaxic Coordinates, Academic Press). Implants are secured by dental cement. A bolus of Lactated Ringers of the 0.9% saline is given at the end of surgery (5mls SC in rats and 1 ml SC in mice after fluids are warmed to normal body temperature) to prevent dehydration. Buprenorphine (0.1-0.5mg/kg SC) is administered twice daily and, then, on an as-needed basis, if animal appears to be in pain. Local antibiotic treatment (bacitracin ointment) and systemic antibiotic treatment (penicillin 100,000 IU/kg IM every 12 hours for the first 48 hours post-op) are administered if post-operative infections occur.
Following surgery, animals are individually housed with food and water available ad libitum. At least one week is allowed for recovery before microdialysis and euthanasia. Following recovery from surgery, the animals are transferred to a microdialysis cage and microdialysis probes are inserted and cemented in the guide shafts that have been installed during surgery. Probe insertion does not cause pain or discomfort because the probe is bypassing skin, muscle and meningeal tissue through the guide shaft. Therefore, probe insertion is done without anesthesia and any anesthesia-induced effects on neurochemistry or behavior are avoided. We let the probes stabilize for 12 hours and then we start sampling every 30 minutes for another 8-12 hours depending on the experiment. We monitor locomotor behavior of the animal through photocells or manual recording of movement by the experimenter. Microdialysate samples are injected into a High Performance Liquid Chromatography with Electrochemical Detection (HPLC-EC) instrument for neurochemical detection and analysis. We look for effects on basal neurochemistry and locomotor behavior. At the end of the experiment the animal is euthanized by an overdose of systemic ketamine (200 mg/kg i.p.) and xylazine (20 mg/kg, i.p.). Then the heart is perfused with 0.9% saline followed by 4% paraformaldehyde. The brains are removed, frozen and cut along the microdialysis probe tract to verify accurate probe placement.
Procedure
<ol>
<li>Set up the stereotaxic instrument and all the materials needed. Make sure the area and instruments are cleaned and sterilized. </li>
<li>Shave off fur with electric razor. Go from the ears to just in-between the eyes, move razor in different directions to effectively clean area of fur. Apply povidine/iodine to shaved area but protect the eyes from it. </li>
<li>Mount the animal onto the stereotaxic apparatus by placing the ear bars into the ear canal and tightening into place. First mount one ear bar in the ear canal, and then hold it in place and slide in the other ear bar. You know you are in the correct location when the head can no longer be moved side to side. Secure the mouth with the anterior mount of the stereotaxic and make sure that the head is level with a ruler. Put the ruler in a vertical position with respect to the stereotaxic instrument platform and check for a 90&deg; angle between the ruler and the middle of the animal&rsquo;s scalp). Confirm this through the stereotaxic instrument if it offers such capability. </li>
<li>Make an anterior/posterior incision on the scalp with a sterile scalpel extending from the lambda to just in-between the eyes of the animal. Use sterilized hemostats to pinch off the skin and keep the incision open. Using several sterile cotton swabs, dry off the exposed skull surface. </li>
<li>Put the guide cannula onto its mount, find bregma on the skull, and position the guide cannula right over this location. Write down the anterior/posterior and lateral coordinates. From bregma, find the correct coordinates needed for the placement of your probe with the aid of the stereotaxic atlas. Position the guide cannula to the correct coordinates by adding or subtracting from bregma. Bring your guide cannula down until it is touching the skull, and then record this ventral coordinate. Make a pencil mark with a sterile pencil at this location on the skull; this is where you will be drilling. </li>
<li>Remove the guide cannula and sterilize your drill bit. Carefully drill a hole at the pencil&nbsp;mark until you get through the width of the skull. Check with the guide cannula to see if it would clear the hole without touching the sides. Keep drilling and checking until the cannula can clear in a straight path. Once the hole is made, use a sterile needle to gently punch the meninges in order to allow unobstructed insertion of the cannula. </li>
<li>Next, using a hand drill, make six holes for skull screws: two anterior to the cannula hole, two lateral to the cannula hole, and two posterior to the sides. Sterilize six screws and place them onto the skull until they are tightly anchored on. </li>
<li>Clean the guide cannula with ethanol and saline, mount, and lower it slowly to the proper ventral coordinate. Make sure that the sides are not touching and that it is going in perfectly vertical. </li>
<li>Place the anchor screw medially and behind the posterior skull screws and hold it in place with tweezers. Mix a thin batch of liquid dental cement and cover the guide cannula, screws, and the rest of the skull with a sterile spatula. Make another batch, thicker this time, and completely cover the area and the cannula and anchor screw enough to secure it. </li>
<li>As the cement becomes thicker and before it solidifies, separate the skin from the cement cup and mold the cement cup with the spatula to make sure the cement cap is smooth all around and does not irritate the skin later. </li>
<li>Allow the dental cement to completely dry before removing the animal from the apparatus. Remove the hemostats. Apply bacitracin all the way around the cement cap. </li>
<li>Once the animal is off the stereotaxic instrument, inject it with 0.25 ml of penicillin IM (intra-muscular) followed by 1 ml of saline SC (subcutaneously). </li>
<li>Place the animal in its own cage and monitor it until it becomes conscious before returning it to its room to recover. </li>
<li>Monitor animals until they recover from anesthesia on the day of surgery and daily post-op, until the end of the experiment, for signs of infection and evaluation of pain/distress. This includes weekends and holidays. Low spontaneous movement, distress vocalization upon handling, hunched posture, diarrhea, swelling and discharge in the area of the headmount, and lack of feeding/drinking are all signs of pain and distress. Buprenorphine (0.1-0.5mg/kg SC) is administered twice daily, and then, on an as-needed basis, if animal appears to be in pain. Local antibiotic treatment (bacitracin ointment) and systemic antibiotic treatment (penicillin 100,000 IU/kg IM every 12 hours for the first 48 hours post-op) are administered if post-operative infections occur. If any of these symptoms persist following administration of buprenorphine, supplemental fluid, and antibiotic treatment within 12 hours of surgery, the animal is euthanized.</li>
</ol>
&nbsp;

<ol>
	<li>Bungay, P. M., Newton-Vinson, P., Isele, W., Garris, P. A., Justice, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microdialysis+of+dopamine+interpreted+with+quantitative+model+incorporating+probe+implantation+trauma.">Microdialysis of dopamine interpreted with quantitative model incorporating probe implantation trauma.</a> J. Neurochem. 86, 932-946 (2003).</li><li>Chen, K. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+tissue+trauma+on+the+characteristics+of+microdialysis+zero-net-flux+method+sampling+neurotransmitters.">Effects of tissue trauma on the characteristics of microdialysis zero-net-flux method sampling neurotransmitters.</a> Journal of Theor. Biology. 238, 863-881 (2006).</li><li>Geiger, B. M., Behr, G. G., Frank, L., Caldera-Siu, A. D., Beinfeld, M. C., Kokkotou, E. G., Pothos, E. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evidence+for+defective+mesolimbic+dopamine+exocytosis+in+obesity-prone+rats.">Evidence for defective mesolimbic dopamine exocytosis in obesity-prone rats.</a> FASEB Journal. 8, 2740-2746 (2008).</li><li>Pothos, E. N., Creese, I., Hoebel, B. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Restricted+eating+with+weight+loss+selectively+decreases+extracellular+dopamine+in+the+nucleus+accumbens+and+alters+dopamine+response+to+amphetamine,+morphine+and+food+intake.">Restricted eating with weight loss selectively decreases extracellular dopamine in the nucleus accumbens and alters dopamine response to amphetamine, morphine and food intake.</a> The Journal of Neuroscience. 15, 6640-6650 (1995).</li></ol>

The authors have nothing to disclose.

In vivo microdialysis is the tool of choice for measuring multiple neurotransmitters and metabolites in distinct brain sites of a living animal. However, it only monitors extracellular levels of neurochemicals and it does not offer the time resolution to monitor neurotransmitter exocytosis in real time. Through a version of the technique called "net-flux", the actual neurotransmitter concentration at a given site can be calculated, which in turn can give accurate measurements of neurotransmitter rate of reuptake through ...

Materials are described in the protocol document

survivable stereotaxic surgery in rodents 

The ability to measure extracellular basal levels of neurotransmitters in the brain of awake animals allows for the determination of effects of different systemic challenges (pharmacological or physiological) to the CNS. For example, one can directly measure how the animal's midbrain dopamine projections respond to dopamine-releasing drugs like d-amphetamine or natural stimuli like food. In this video, we show you how to implant guide cannulas targeting specific sites in the rat brain, how to insert and implant a microdialysis probe and how to use high performance liquid chromatography coupled with electrochemical detection (HPLC-EC) to measure extracellular levels of oxidizable neurotransmitters and metabolites. Local precise introduction of drugs through the microdialysis probe allows for refined work on site specificity in a compound s mechanism of action. This technique has excellent anatomical and chemical resolution but only modest time resolution as microdialysis samples are usually processed every 20-30 minutes to ensure detectable neurotransmitter levels. Complementary ex vivo tools (i.e., slice and cell culture electrophysiology) can assist with monitoring real-time neurotransmission.

Watch this Scientific Journal Video about Survivable Stereotaxic Surgery in Rodents at JoVE.com

Survivable Stereotaxic Surgery in Rodents

The monitoring of extracellular neurotransmitter levels in distinct brain regions of freely moving animals offers insights on the link between neurotransmitter release and behavior. In vivo microdialysis coupled with electrochemical detection provides excellent anatomical and chemical resolution; and information on how basal neurotransmission is altered by pharmacological or physiological manipulations.

Survivable Stereotaxic Surgery in Rodents 

The ability to measure extracellular basal levels of neurotransmitters in the brain of awake animals allows for the determination of effects of different ...

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68.1K Views.  Tufts University. The monitoring of extracellular neurotransmitter levels in distinct brain regions of freely moving animals offers insights on the link between neurotransmitter release and behavior. In vivo microdialysis coupled with electrochemical detection provides excellent anatomical and chemical resolution; and information on how basal neurotransmission is altered by pharmacological or physiological manipulations. 

Video: Survivable Stereotaxic Surgery in Rodents

A Craniotomy Surgery Procedure for Chronic Brain Imaging

Imaging techniques are becoming increasingly important in the study brain function. Among them, two-photon laser scanning microscopy has emerged as an extremely useful method, because it allows the study of the live intact brain. With appropriate preparations, this technique allows the observation of the same cortical area chronically, from minutes to months. In this video, we show a preparation for chronic in vivo imaging of the brain using two-photon microscopy. This technique was initially pioneered by Dr. Karel Svoboda, who is now a Howard Hughes Medical Institute Investigator at Janelia Farm. Preparations like the one shown here can be used for imaging of neocortical structure (e.g., dendritic and axonal dynamics),  to record neuronal activity using calcium-sensitive dyes, to image cortical blood flow dynamics, or for intrinsic optical imaging studies. Deep imaging of the neocortex is possible with optimal cranial window surgeries. Operating under the most sterile conditions possible to avoid infections, together with using extreme care to do not damage the dura mater during the surgery, will result in successful and long-lasting glass-covered cranial windows.


This video and protocol demonstrate how to implant a glass-covered cranial window in rodents. These preparations can be used for chronic in vivo two-photon imaging of the neocortex over time scales of months. It may also be used for other types of imaging, including optical intrinsic signal imaging.

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Dissecting the Non-human Primate Brain in Stereotaxic Space

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The non-human primate is an important translational species for our understanding of the normal processing of the brain. The anatomical organization of the primate brain can provide important insights into normal and pathological conditions in humans.

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Establishing Primary Adult Fibroblast Cultures From Rodents

The importance of using primary cells, rather than cancer cell lines, for biological studies is becoming widely recognized. Primary cells are preferred in studies of cell cycle control, apoptosis, and DNA repair, as cancer cells carry mutations in genes involved in these processes. Primary cells cannot be cultured indefinitely due to the onset of replicative senescence or aneuploidization. Hence, new cultures need to be established regularly. The procedure for isolation of rodent embryonic fibroblasts is well established, but isolating adult fibroblast cultures often presents a challenge. Adult rodent fibroblasts isolated from mouse models of human disease may be a preferred control when comparing them to fibroblasts from human patients. Furthermore, adult fibroblasts are the only available material when working with wild rodents where pregnant females cannot be easily obtained. Here we provide a protocol for isolation and culture of adult fibroblasts from rodent skin and lungs. We used this procedure successfully to isolate fibroblasts from over twenty rodent species from laboratory mice and rats to wild rodents such as beaver, porcupine, and squirrel.

This article describes a protocol for isolation and maintenance of primary fibroblast cultures from skin and lung tissue of wild rodents.

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Principles of Rodent Surgery for the New Surgeon

For both scientific and animal welfare reasons, training in basic surgical concepts and techniques should be undertaken before ever seeking to perform surgery on a rodent. Students, post-doctoral scholars, and others interested in performing surgery on rodents as part of a research protocol may not have had formal surgical training as part of their required coursework. Surgery itself is a technical skill, and one that will improve with practice. The principles of aseptic technique, however, often remain unexplained or untaught. For most new surgeons, this vital information is presented in piecemeal fashion or learned on the job, neither of which is ideal. It may also make learning how to perform a particular surgery difficult, as the new surgeon is learning both a surgical technique and the principles of asepsis at the same time. This article summarizes and makes recommendations for basic surgical skills and techniques necessary for successful rodent surgery. This article is designed to supplement hands-on training by the user's institution.

Before attempting surgery, a new surgeon should have training in basic surgical techniques and concepts. This article will present basic surgical considerations with an emphasis on rodents.

For both scientific and animal welfare reasons, training in basic surgical concepts and techniques should be undertaken before ever seeking to perform ...

Intracavernosal Pressure Recording to Evaluate Erectile Function in Rodents

Erectile dysfunction (ED) is defined as the inability to attain or keep an erection of the penis, and this has become a prevalent male sexual disorder. Rodents are employed by many studies to research the physiology/pathology of erectile function. Erectile function in rodents can be evaluated by measuring the intracavernosal pressure (ICP). In practice, ICP can be monitored following electrical stimulation of the cavernous nerves (CNs). The arterial pressure of the carotid artery (the mean arterial pressure) is used as the reference for ICP. Using ICP recording protocols, many key parameters of erectile function can be measured from the ICP response curve. The ICP measurement provides more information than the apomorphine-induced penile erection test, and is cheaper than telemetric monitoring of the corpus spongiosum penis, making this method the most popular one to evaluate erectile function. However, compared to the easily-performed APO-induced erectile function test, successful ICP recordings require attention to detail, practice, and adherence to the operation method. In this work, an introduction to ICP recording in rats is provided to complement the procedure efficiently.

Intracavernosal pressure recording (ICP) is an important method to evaluate the erectile function of experimental animals. Here, a detailed protocol is demonstrated for the recording procedure of ICP by catheterizing the crura penis and then electrically stimulating the cavernous nerves in rats.

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Performing In Vivo and Ex Vivo Electrical Impedance Myography in Rodents

Electrical impedance myography (EIM) is a convenient technique that can be used in preclinical and clinical studies to assess muscle tissue health and disease. EIM is obtained by applying a low-intensity, directionally focused, electrical current to a muscle of interest across a range of frequencies (i.e., from 1 kHz to 10 MHz) and recording the resulting voltages. From these, several standard impedance components, including the reactance, resistance, and phase, are obtained. When performing ex vivo measurements on excised muscle, the inherent passive electrical properties of the tissue, namely the conductivity and relative permittivity, can also be calculated. EIM has been used extensively in animals and humans to diagnose and track muscle alterations in a variety of diseases, in relation to simple disuse atrophy, or as a measure of therapeutic intervention. Clinically, EIM offers the potential to track disease progression over time and to assess the impact of therapeutic interventions, thus offering the opportunity to shorten the clinical trial duration and reduce sample size requirements. Because it can be performed noninvasively or minimally invasively in living animal models as well as humans, EIM offers the potential to serve as a novel translational tool enabling both preclinical and clinical development. This article provides step-by-step instructions on how to perform in vivo and ex vivo EIM measurements in mice and rats, including approaches to adapt the techniques to specific conditions, such as for use in pups or obese animals.

Performing In Vivo and Ex Vivo Electrical Impedance Myography in Rodents

This article details how to perform in vivo (using surface and needle electrode arrays) and ex vivo (using a dielectric cell) electrical impedance myography on the rodent gastrocnemius muscle. It will demonstrate the technique in both mice and rats and detail the modifications available, (i.e., obese animals, pups).

Electrical impedance myography (EIM) is a convenient technique that can be used in preclinical and clinical studies to assess muscle tissue health and ...

Quantification of Skeletal Muscle Fatty Infiltration via Decellularization

Decellularization-Based Quantification of Skeletal Muscle Fatty Infiltration

Fatty infiltration is the accumulation of adipocytes between myofibers in skeletal muscle and is a prominent feature of many myopathies, metabolic disorders, and dystrophies. Clinically in human populations, fatty infiltration is assessed using noninvasive methods, including computed tomography (CT), magnetic resonance imaging (MRI), and ultrasound (US). Although some studies have used CT or MRI to quantify fatty infiltration in mouse muscle, costs and insufficient spatial resolution remain challenging. Other small animal methods utilize histology to visualize individual adipocytes; however, this methodology suffers from sampling bias in heterogeneous pathology. This protocol describes the methodology to qualitatively view and quantitatively measure fatty infiltration comprehensively throughout intact mouse muscle and at the level of individual adipocytes using decellularization. The protocol is not limited to specific muscles or specific species and can be extended to human biopsy. Additionally, gross qualitative and quantitative assessments can be made with standard laboratory equipment for little cost, making this procedure more accessible across research laboratories.

Author Spotlight: Decellularization-Based Quantification of Skeletal Muscle Fatty Infiltration  

The present study describes decellularization-based methodologies for visualizing and quantifying intramuscular adipose tissue (IMAT) deposition through intact muscle volume, as well as quantifying metrics of individual adipocytes that comprise IMAT.

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Rearing Germ-Free &lt;em&gt;Nasonia&lt;/em&gt; Wasps

Optimizing the Rearing Procedure of Germ-Free Wasps

Aseptic rearing technology is a method of culturing insects under sterile or almost sterile conditions, which can effectively eliminate the influence of external microorganisms on insect microbiota and thus promote the rapid development of insect microbiota research. Nasonia (wasp genus) is a parasitic wasp insect that has many advantages, such as a short lifespan, high genetic variation, easy operation, etc., and is widely used as an insect model system. Unlike antibiotic treatment, which can only reduce the number of microorganisms in animals, aseptic rearing techniques can control both the composition and quantity of microorganisms in animals, further facilitating the study of host-microbe interactions. However, previous versions of Nasonia rearing medium (NRM) have some defects and problems, such as a complex and time-consuming preparation process, easy contamination by bacteria or fungi, and short storage time. Therefore, this study solves these problems by optimizing the tools used in the NRM preparation process, storage conditions, and component ratios. The optimized medium could allow storage at -20 &#176;C for at least 3 months and eliminate the possibility of NRM contamination during feeding sterile wasps. This further improves the survival rate and health level of aseptic Nasonia, which is important for using Nasonia as a model for microbial research.

Author Spotlight: Optimizing the Rearing Procedure of Germ-Free Wasps  

Nasonia wasp embryos were dissected from Lucillia sericata pupae after parasitization for 12-24 h and washed with alcohol and 10% sodium hypochlorite solution to obtain germ-free embryos. After rearing the germ-free embryos&#160;and supplying them with Nasonia rearing medium to grow and develop in vitro, germ-free Nasonia adults were obtained.

Aseptic rearing technology is a method of culturing insects under sterile or almost sterile conditions, which can effectively eliminate the influence of ...

Replicating Cataract Surgical Techniques in a Mouse Model Through LEC Extraction

Modeling Cataract Surgery in Mice

Cataract surgery (CS) is an effective treatment for cataracts, a major cause of visual disability worldwide. However, CS leads to ocular inflammation, and in the long term, it can result in posterior capsular opacification (PCO) and/or lens dislocation driven by the post-surgical overgrowth of lens epithelial cells (LECs) and their conversion to myofibroblasts and/or aberrant fiber cells. However, the molecular mechanisms by which CS results in inflammation and PCO are still obscure because most in vitro models do not recapitulate the wound healing response of LECs seen in vivo, while traditional animal models of cataract surgery, such as rabbits, do not allow the genetic manipulation of gene expression to test mechanisms. Recently, our laboratory and others have successfully used genetically modified mice to study the molecular mechanisms that drive the induction of proinflammatory signaling and LEC epithelial to mesenchymal transition, leading to new insight into PCO pathogenesis. Here, we report the established protocol for modeling cataract surgery in mice, which allows for robust transcriptional profiling of the response of LECs to lens fiber cell removal via RNAseq, the evaluation of protein expression by semi-quantitative immunofluorescence, and the use of modern mouse genetics tools to test the function of genes that are hypothesized to participate in the pathogenesis of acute sequelae like inflammation as well as the later conversion of LECs to myofibroblasts and/or aberrant lens fiber cells.

Author Spotlight: Unraveling the Molecular Mechanisms in PCO and Fibrosis Following Cataract Surgery

This method models cataract surgery in vivo by removing lens fiber cells from adult mice and leaving behind the capsular bag with attached lens epithelial cells (LECs). The injury response is then assessed at various times post-surgery using molecular and morphological criteria.

Cataract surgery (CS) is an effective treatment for cataracts, a major cause of visual disability worldwide. However, CS leads to ocular inflammation, and ...

Sagittal Suture Expansion Surgery to Prepare Mouse Suture Expansion Model

Begin by preparing two retention holders. Use light wire pliers and a 0.014 inch Australian wire to make a helical loop. Then prepare customized stainless steel springs and cut the spring into an appropriate length.Also prepare one straight Australian wire or straight steel wire of length seven millimeters and cut two pieces of paper with diameters of two millimeters to use as barriers. Next, place an anesthetized mouse on the operating table and carefully remove the fur on top of the head with hair removal cream or a shaver. Then disinfect the scalp of the mouse using alternating rounds of Iodoform and 75%alcohol.Position the mouse with its head on the top and use surgical tapes to fix the limbs on the operating table. Now, use surgical scissors to perform an arcuate flap along the scalp near the neck of the mouse, fully exposing the sagittal suture and surrounding skull. Then, fix the scalp flap with a 6-0 suture on the operating table.Dry the skull and etch it with 37%phosphoric acid for 20 seconds. Then use normal saline to clean up the acid etching. Place the two holders on both sides of the parietal bones, three millimeters from the sagittal sutures and glue them with a light cured adhesive.Then put back the scalp flap. Label the position of the holders and cut two small holes in the scalp at the corresponding positions. Reset the flap scalp by simultaneously passing the small loops through the holes to expose the skin surface.To install the spring and guide wire, cut the spring one millimeter longer than the distance between the two holders. Compress the pressure spring and place it between the small coils on both sides. Pass the stainless steel wire through the small coils and the spring and release the spring to obtain a starting thrust of about 30G.Next, place two scraps of paper between the spring and the holders and bond them with a light cured adhesive to set up barriers at both ends of the spring. After expansion, micro CT was used to visualize the changes in the cranial suture. In comparison to controls, the cranial suture gradually and significantly expanded after applying force for one day and fluffy bony protrusions appeared on the bone edge after day five.Also, visualization of mineralization using double labeling method, showed that force loading significantly activated osteogenesis. Newly mineralized bone was observed after seven days, indicating the bone remodeling process after suture expansion.