Two-Photon Intravital Microscopy of Glioblastoma in a Murine Model

Summary

We present a novel approach for two-photon microscopy of the tumor delivery of fluorescent-labeled iron oxide nanoparticles to glioblastoma in a mouse model.

Abstract

The delivery of intravenously administered cancer therapeutics to brain tumors is limited by the blood-brain barrier. A method to directly image the accumulation and distribution of macromolecules in brain tumors in vivo would greatly enhance our ability to understand and optimize drug delivery in preclinical models. This protocol describes a method for real-time in vivo tracking of intravenously administered fluorescent-labeled nanoparticles with two-photon intravital microscopy (2P-IVM) in a mouse model of glioblastoma (GBM).

The protocol contains a multi-step description of the procedure, including anesthesia and analgesia of experimental animals, creating a cranial window, GBM cell implantation, placing a head bar, conducting 2P-IVM studies, and post-surgical care for long-term follow-up studies. We show representative 2P-IVM imaging sessions and image analysis, examine the advantages and disadvantages of this technology, and discuss potential applications.

This method can be easily modified and adapted for different research questions in the field of in vivo preclinical brain imaging.

Introduction

Two-photon intravital microscopy (2P-IVM) is a fluorescence imaging technique that allows the visualization of living tissue¹.

First developed in the 1990s, 2P-IVM has been used for in vivo analysis of the retina², kidney³, small intestine⁴, cochlea⁵, heart⁶, trachea^7, and the brain in various preclinical models^8,9. In the field of neuroscience, 2P-IVM has gained importance as a technique for real-time imaging of the healthy brain in awake animals¹⁰, as well as studying diseases of the nervous system such as Alzheimer's¹¹, Parkinson's¹² and glioblastoma (GBM)^13,14,15,16.

2P-IVM offers an elegant solution for studying the tumor microenvironment during the development of GBM. While some previous studies focused on in vitro¹⁷ and ex vivo models¹⁸, others implemented orthotopic¹⁹ and xenotropic²⁰ in vivo models for examining GBM. Madden et al. performed native imaging of CNS-1 rat glioma cell line in a mouse model¹³. Using an orthotopic GL261-DsRed murine model, Ricard et al. performed an intravenous administration of a fluorophore to enhance the blood vessels in the tumor region in 2P-IVM¹⁴.

Here, we apply 2P-IVM for tracking the tumor delivery of fluorescent-labeled iron oxide nanoparticles (NP) in an orthotopic mouse model of GBM. Using a cranial window, this method allows us to study the real-time spatiotemporal distribution of NPs in the brain in detail.

Protocol

The animal procedure described in this protocol is in accordance with the requirements of the Administrative Panel on Laboratory Animal Care (APLAC).

1. Cell culture

1. Preparation of hood
   1. Wash hands, wear gloves and a lab coat. Turn on the biological safety cabinet and set the sash level to an appropriate opening height. Let the hood purge for 3-5 min. Spray the hood area with 70% ethanol and wipe it down with tissue paper.
   2. Spray all reagents with 70% ethanol and wipe down with tissue paper. Move the reagents into the hood. Do not put any items on the grill. If a spill occurs in the hood, cover it with wipes, spray it with 70% ethanol, and wipe again.
   3. After the experiment, carefully close the lids of each item, wipe down all materials, remove any items from the hood, and transfer them to their original location. Spray 70% ethanol in the hood and wipe it down. Shut the sash and turn off the biological safety cabinet.
2. Preparation of a growth medium
   1. Add 50 mL of 100% fetal bovine serum (FBS) to 500 mL Dulbecco’s Modified Eagle Medium (DMEM). Add 5 mL of the antibiotic-antimycotic solution (100x).
   2. Pass all the reagents through a sterile 500 mL filtering bottle (0.22 µm filter). 
3. Thawing of cryopreserved cells
   1. In a laminar flow hood, add 20 mL of the growth medium to a T75 flask. Label the flask with the name of cells, date, and passage number on the flask. In this study, adherent C6 glioma cells were used.
   2. Thaw cells quickly in a 37 °C water bath. Do not vortex the cells. Thaw and immediately use the cells.
   3. Transfer the thawed cells to the T75 flask using a 1 mL pipette tip. Be careful not to introduce any bubbles during the transfer process, and avoid any medium residue in the neck of the flask. This could increase the risk of possible contamination.
4. Changing the medium
   1. Change the medium on the day following thawing to remove any residual trypsin or dimethyl sulfoxide (DMSO) (step 1.6). Afterward, change the medium at least two times per week or more, depending on cell confluency level. The medium must be changed one day after passage to eliminate trypsin.
   2. To change the medium, turn the aspiration apparatus on, attach the aspiration glass pipet, and aspirate all the medium.
   3. Add 20 mL of the growth medium (step 1.2).
5. Passaging the cells
   1. Load the hood with the items needed in the course of the experiment.
   2. Use an aspiration glass pipet and aspirate away all media, ensuring the glass pipette is on the non-cell side of the flask. For this step, position the T75 flask vertically.
   3. Add ~10 mL of room temperature (RT) phosphate-buffered saline (PBS, Ca⁺⁺ and Mg⁺⁺ free) or Hanks' Balanced Salt Solution (HBSS, Ca^++, and Mg⁺⁺ free) on the non-cell side. Wash cells briefly with PBS by positioning the T75 flask horizontally with the cell side facing down.
   4. Immediately place the flask vertically and aspirate the PBS/HBSS. Repeat this washing and aspirating 3 times.
   5. Add 2 mL of trypsin (recombinant or animal-derived) on the cell side (T75 flask horizontal, cell side facing down). Incubate in a standard tissue culture incubator (37 °C, 5% CO₂) for 4 min. Triturate it once after 2 min using a 5 mL pipette.
   6. After 4 min of total incubation, transfer the solution to a 15 mL conical tube. Add 8 mL of the growth solution to the conical tube (2 mL trypsinized cells + 8 mL of medium = 10 mL in total).
   7. Use another empty 15 mL conical tube with a filter and transfer the cells via a 25 mL pipette to achieve individual cells.
   8. Transfer 1/10 (1 mL) of the solution (cell + media + Tryp-LE) to a T75 flask with 20 mL of media. Alternatively, count the cells (step 1.7). Change the media the next day to have a non-trypsin media solution.
6. Freezing the medium and the cells
   1. Thaw 100% FBS in a 4 °C refrigerator. Do not use a water bath or heat since this may damage the proteins in FBS.
   2. To prepare 50 mL of freezing media, mix 45 mL of DMEM, 5 mL of FBS, and 5 mL of DMSO. Aliquot it to 1-2 mL containers to prevent intermittent thawing and protein damage. Store them in -20 ° or -80 °C freezer.
   3. For the remaining 9 mL of solution (step 1.5), add 1 mL of medium to reach 10 mL in total. Centrifuge at 300 x g for 4 min.
   4. Remove supernatant and resuspend in 1 mL of freezing medium (see above). Place the cells in a freezing container in -80 °C freezer. Move the cells to liquid N₂ for long-term storage after 1 or 2 days.
7. Counting the cells
   1. For the remaining 9 mL (step 1.5), add 1 mL of media to reach 10 mL in total. Centrifuge at 300 x g for 4 min.
   2. Remove the supernatant and resuspend in 4 mL of HBSS. Resuspend 10 µL of cells in 80 µL of HBSS + 10 µL of 0.4% Trypan Blue, dilution factor = 10.
   3. Take 17 µL of the solution and count four 4 x 4 squares in a hemocytometer. Average counts for the four 4 x 4 squares and multiply by 10⁴x dilution factor to get cells/mL.

2. Surgery

NOTE: It is recommended to perform the surgery by two researchers, where one person is responsible for preparing the cells, mixing the dental cement, and generally assisting in the procedure, while the second person focuses on remaining sterile. Having a second manipulator to assist with the surgical procedure considerably reduces the likelihood of contamination occurring. Following best surgical practices would reduce the chances of post-operative complications. Figure 1A provides an overview of the components of the cranial window.

1. General preparations
   1. Confirm that the procedure is approved by the local animal welfare institutional guidelines before starting.
      NOTE: This study used 5-month-old female NSG mice (NOD.Cg-Prkdcscid Il2rgtm1Wjl/SzJ) (n = 5).
   2. Before surgery, autoclave all surgical instruments and ensure that all necessary supplies are available. Print the anesthesia and surgery records.
   3. Ensure that upon arrival, the animals have at least 1 week of acclimatization to the animal husbandry room to reduce additional post-operative distress.
   4. On the day of surgery, ensure that all devices (microscope, heating pad, sterilizer, drill, vacuum, etc.) are ready to use. Confirm that the anesthesia machine contains enough isoflurane and refill if necessary. For new users, the cranial window procedure can take up to 2 h per animal; therefore, having enough isoflurane is crucial.
   5. Place the glass windows and head bars in alcohol and prepare a container with saline. This will ensure a smooth workflow without any major breaches in sterility and keep the time under anesthesia for each animal as short as possible.
   6. Open all surgical materials on a sterile surface on the working station. For example, the inside of a sterile gauze packaging can be used for this purpose. Use the tips-only technique. When not using surgical instruments, place the tips on a sterile surface, such as the inside of the packaging of sterile gauze.
   7. When performing multiple surgeries, place all surgical instruments in the sterilizer in between surgeries to disinfect them. When performing surgeries on more than 5 animals, use a new set of autoclaved instruments. Switch out the drill tip when it becomes blunt for a new one to avoid tissue trauma.
2. Anesthesia and preparations for surgery
   NOTE: The anesthesia setup is identical for the surgery, as well as for imaging.
   1. Anesthetize the mouse using isoflurane (3%-5% for induction) in an anesthetic chamber. After ensuring adequate anesthetic depth by testing the pedal withdrawal reflex (foot pad pinch on both hind feet), transfer the animal to a preparation working station. Maintain anesthesia over a nose cone (1%-2%). Here, apply an eye ointment to prevent corneal damage. Regularly control the loss of reflexes during surgery by checking the paw reflex.
   2. If any identification of the animal is required, use an ear punch tool or scissors to mark the ears or use a marker to mark the tail.
   3. Remove the fur covering the skull between the ears and eyes with a depilatory cream. Leave the cream as long as indicated (30-60 s) by the manufacturer to avoid skin irritation. Make sure that the cream comes in contact with the hair roots by applying it against the direction of hair growth. Avoid that the depilatory cream comes in contact with the eyes. Remove any cream and hair remnants by cleaning the area with saline. Alternatively, use clippers.
   4. To avoid any intra- and post-operative pain, infection, or swelling, administer non-steroidal anti-inflammatory drugs (NSAIDs), opioids, anti-inflammatory agents, and antibiotics. Administer carprofen (5-20 mg/kg, subcutaneous [SQ]), buprenorphine - sustained release (1 mg/kg SQ), cefazolin (20 mg/kg SQ), and dexamethasone (0.2 mg/kg SQ) before surgery. Administer approximately 0.3 mL of 0.9% saline SQ to avoid dehydration.
   5. Following, scrub the area by alternately swabbing povidone-iodine and isopropyl alcohol three times in circular motions from the middle of the skull to the periphery.
3. Craniotomy procedure
   1. Transfer the animal to the surgical table and place it in ventral recumbency on a heating pad (~37 °C) to ensure isothermal conditions during the procedure. Hypothermia in rodents significantly reduces survival rates and prolongs the post-operative recovery phase.
   2. Place the head in the stereotactic frame by positioning the ear bars and incisor bars. For the ear bars, find the zygomatic arch and insert the bars behind the arches. Start by securing the contralateral bar with the dominant hand (e.g., secure the left bar with your right hand if you are right-handed) and then secure the opposite side. Adjust the positions of the ear and incisor bars by turning the screws if required.
   3. Reapply eye ointment if necessary, and use a piece of aluminum foil to cover the eyes. This will prevent any damage caused by the bright microscope light and UV light used to cure the cyanoacrylate glue later (step 2.5).
   4. Before incision, check again for toe pinch reflex; if necessary, adjust anesthesia accordingly. Connect the animal to the anesthesia monitoring device by positioning the sensor on the hind paw. Measuring the heart rate and blood oxygen concentration would help reduce mortality and improve post-operative recovery. Additionally, acquire the respiratory rate by visually inspecting the chest movement.
   5. Cover the body of the animal with a drape to avoid hypothermia and contamination. Perform one final povidone-iodine swab before starting the surgery.
   6. Put on gloves and a clean lab coat.
      NOTE: Using sterile gloves is encouraged due to the invasiveness of the procedure and to lower the risk of any post-operative infection and inflammation resulting in morbidity and loss of image quality in 2P-IVM (section 5).
   7. Lift the skin on the skull and create an incision by holding the scissors between the right eye and ear and cutting towards the left side of the skull following the incision marks. Remove the resulting round skin flap.
      NOTE: Depending on the region of interest in the brain and the window size, the incision site and diameter can vary. The size of the incision needs to be bigger than the diameter of the head to accommodate room for mounting (step 2.5). If needed, the skin surrounding the incision can be gently dissected to create more space.
   8. Remove the periosteum by gently scratching the skull surface with a scalpel blade or a microcurrette. Exercise caution when removing the periosteum around the cranial sutures since those areas are fragile and more prone to bleeding. Use saline and a vacuum to keep the surgical area clean from debris. Scratch the periosteum to ensure better adherence to the cyanoacrylate glue and dental cement (step 2.5)
   9. Identify the region of the brain to perform the cell injection. For this, use an atlas of the stereotactic coordinates of the mouse brain. Per the brain coordinates, mark the position of the cranial window by using a biopsy punch in the same diameter as the window, a surgical pen, or an autoclaved pencil.
   10. Hold the drill in the dominant hand and any other instrument (syringe, forceps) in the non-dominant hand. Avoid prolonged drilling in the same spot. The drill should be used in bursts to avoid overheating. During these breaks, apply cold saline to the calvarium to keep the surgical view clean and to prevent overheating. Use the sensory feedback from the drill tip to detect when the skull has been perforated completely.
   11. Use cold saline, in combination with a vacuum, to clean the skull surface. Do not use gauze or a cotton tip applicator after opening the calvarium since remnants of cotton threads may lead to a foreign body reaction upon sealing the brain window.
   12. While drilling, ensure that the trajectory and diameter are the same size as those of the glass coverslip. Do this by positioning the glass coverslip on top of the skull and confirming that the glass coverslip will fit exactly inside the cranial window.
   13. Once it is possible to depress the skull flap by gently pressing on it, use forceps to carefully remove the fragment from the surgical field (Figure 1B, left). If it is not yet possible to depress the skull, continue drilling in the areas of the cranial window that are still connected to the rest of the skull and reevaluate.
   14. If minor bleeding occurs, use a hemostatic sponge combined with light pressure or alternatively, ice cold saline, to help reduce blood loss.
4. Cell implantation
   1. Prepare and count the cells to be implanted as described in (step 1.7). Ensure that the cell number is 200,000 cells/µL.
      NOTE: Suspending the cells in a membrane matrix that solidifies at RT is recommended. This will improve the success rate of implanted cells in the brain parenchyma.
   2. Transfer the cells in a container with ice to the surgical room. Use a gastight microliter syringe. Before aspiration, keep the syringe on ice to avoid temperature differences.
   3. After aspirating 1 µL, position the syringe inside the stereotactic frame.
      NOTE: An automated stereotactic coordinate device showing the positions in the X, Y, and Z axes can be used to save time. It is also possible to manually calculate the positions based on lambda. Injecting more than 1.5 µL is not recommended. This will ensure that the injected area does not overfill, causing unsuccessful implantation, growth outside the brain parenchyma, or metastases.
   4. Perform implantation in the V2MM region (visual cortex 2, mediomedial part) of the brain, which corresponds to approximately medial to lateral (M-L): -1.2 to -1.6 mm, and dorsal to ventral (D-V): -2.6 to -3.5 mm coordinates.
      1. For two-photon imaging (step 4.6), implant the cells in the superficial brain areas so that the tumor mass can be visualized through the cranial window later on. Inject 0.5 µL (100,000 cells) at anterior to posterior (A-P): 0.8 mm depth. Move the needle slowly, in a stepwise manner, when entering the brain and wait for 30 s after injection (Figure 1B, middle column).
      2. Use the same approach when retracting the needle and exiting the brain tissue. Avoid injecting close to the ventricles since the cerebrospinal fluid might encourage metastasis in the central and peripheral nervous systems.
5. Closure of the cranial window
   1. Move the head bar and glass coverslip from alcohol to a saline solution. Use a vacuum tip or forceps to navigate those components to the surgical site.
   2. Position the glass coverslip inside the cranial window and place a small amount of cyanoacrylate glue between the skull and the glass coverslip. UV-activated cyanoacrylate glue reduces the curing time and avoids accidental displacement. If any glue spills on the glass coverslip, remove it with a cotton tip applicator or by gently scratching it off with the blunt side of a scalpel before it cures completely.
      CAUTION: UV light is harmful to the eyes and skin. Avoid eye and skin contact while curing the glue.
   3. After securing the glass coverslip in the cranial window, apply the head bar. Mix the two-component dental cement, position the head bar, and apply the cement to the surrounding area, ensuring contact between the skull and the head bar. Carefully clean any superfluous cement with the tip of a syringe, a scalpel, or a drill tip.
      NOTE: Dental cement solidifies very quickly, so it is advised to apply it in a timely manner.
   4. After sealing the cranial window, identify if any skull areas between the dental cement and skin are still exposed. If that is the case, apply surgical glue to cover up the skull by closing the skin. Alternatively, perform a single suture with an absorbable suture material.

3. Post-surgical recovery and tumor growth

1. Recovery
   1. Monitor the animal on a heated pad in a recovery cage until it regains full consciousness. House the animals separately post-surgery to reduce the risk of injury.
   2. Administer a recovery diet, such as commercially available water gels with electrolytes or sugars. Alternatively, supply moistened standard lab chow to ensure easy nutrition and avoid dehydration and hypoglycemia.
2. Monitoring
   1. Upon recovery, monitor the animal daily for signs of pain, distress, or infection. If needed, administer analgesics, anti-inflammatory drugs, or antibiotics. If an animal reaches the study endpoint, euthanize it humanely. Following the final imaging session, euthanize the mouse by carbon dioxide asphyxiation, followed by cervical dislocation.
      NOTE: Examples of early euthanasia criteria include but are not limited to significant weight loss (>20%), neurological disorders, dyspnea, excessive bleeding during surgery, or pronounced stereotypical behavior. Inspect the cranial window and clean it with saline or alcohol when necessary.
   2. To track tumor growth and plan the imaging time points, perform whole-body bioluminescence imaging (BLI). For this, inject 150 mg/kg D-luciferin intraperitoneally and image the animal after 5-15 min.

4. Nanoparticle synthesis

1. Particle preparation
   1. Add 11.17 mL of Ferumoxytol (6 mmol Fe in total) to a solution containing 50 mL of 5 M NaOH, 20 mL of deionized water (DI water) and 20 mL of epichlorohydrin. Observe phase segregation at this stage. Incubate the mixture at RT under gentle shaking for 24 h.
   2. After 24 h, the solution becomes uniform. Remove excess epichlorohydrin using dialysis tubing (12,000-14,000 Da cutoff) against water for 3 days.
   3. After dialysis, transfer the solution remaining in the tube into a glass bottle (total volume ~110 mL). Add 30 mL of ammonia hydroxide, and stir the mixture at 37 °C for 18-20 h.
   4. After stirring, repeat dialysis against water for 3 days. Transfer the solution remaining in the dialysis tubing into a new glass bottle with a total volume ~110 mL.
2. Fluorescein isothiocyanate (FITC) conjugation
   1. Take out 27.5 mL of amine-functionalized particles for FITC conjugation. Concentrate the particles using a 10 kDa ultracentrifugation filter and wash with a pH 9 (Na₂CO₃/NaHCO₃) buffer at 5752 x g at 25 °C for 10 min.
   2. Add 4 mL of the solution into the filter and ultracentrifuge the tube at 5752 x g at 25 °C for 10 min. Discard the filtrate and refill the filter with buffer for a second round of washing with the same protocol.
   3. Discard the filtrate and collect the solution in the filter using a pipet. Estimate the molar concentration of the particle by the mass concentration of Ferumoxytol, assuming the diameter of each particle is 5 nm.
   4. Dissolve FITC in anhydrous dimethyl sulfoxide (DMSO) at 10 mg/mL. Slowly add 1.4 mL of DMSO-dissolved FITC into the concentrated amine-functionalized particle. The molar ratio of FITC: particle is 20:1. After that, incubate the solution at 4 °C for 8 h in the dark.
   5. Quench the reaction by adding excess NH₃.H₂O (the final concentration of NH₃.H₂O is 50 mM), then letting the solution stay at 4 °C in the dark for 2 h. Wash away excess FITC using Na₂CO₃/NaHCO₃ (pH 9) buffer through a 10 kDa filter at 5752 x g at 25 °C for 10 min. The final pH of the solution is adjusted to 7.4 using HCl aqueous solution.

5. 2P-IVM

NOTE: For the 2P-IVM sessions, a Prairie Ultima IV microscope with a custom stage (Figure 2 and Supplemental Coding File 1) that allows adjusting the position of the cranial window horizontally and vertically was used. Fiji software was used for post-processing and image analysis. This way, the laser beam can be adjusted to hit the glass at a 90 ° angle, reducing artifacts and improving imaging quality.

1. Imaging session
   1. Turn on the Ti-Sapphire laser. Turn on the main system switch and Prairie View software.
   2. Anesthetize the animal, as described above (step 2.2). Perform imaging sessions for up to 2 h per animal. Keep the imaging time to a minimum to reduce the risk of anesthesia complications and associated morbidity or mortality.
   3. Immobilize the mouse under the two-photon microscope using a custom stage (Figure 1B, right column). Place the animal in a prone position on a heated pad designed for rodent surgery and secure it lightly using tape while paying attention to not constrict the thorax. Place a drop of water on the cranial window and adjust the focus.
   4. Acquire heart rate and blood oxygenation to monitor the animals' vital parameters. Position the sensor on the hind paw and keep the monitoring device outside the two-photon imaging chamber.
   5. Once the animal is on the microscope stage, adjust the head position. Using the binoculars and widefield fluorescent illumination, set the red fluorescent protein (RFP) filter and find the desired imaging field of view (Figure 1B, right column).
   6. Once sample orientation and field of view are determined, switch the microscope from widefield mode to light scanning microscopy mode (LSM).
   7. Make sure the Ti-Sapphire laser is mode-locked at 920 nm, and all shutters are open. Bring the GaAsP photomultiplier tube detectors (PMT) voltages up to 500-600 V. Use two PMTs with filter sets with band passes at 595/50 (RFP) and 525/50.
   8. Press Live Image and slowly modify the pockel cell to increase the laser power at the sample until an image is visible.
   9. Once a clear image is achieved, use the software and the motorized stage to set the top and bottom of an image stack within the desired sample area. Beware of the step size and take into consideration that the Z-axis adjusts the depth of the lens.
   10. With increasing depth, the images will get darker. Increase the pockels/PMT gain slowly to keep the image bright. Be careful not to use too much power as it can lead to phototoxicity and tissue damage.
   11. Set a save path and start the Z series. It will automatically traverse to the starting and ending positions using the set pockel/PMT settings. Once the stack is complete, use the playback to look over the stack for quality. Once acquiring the necessary images is complete, turn off the Ti-Sapphire laser.
   12. Move the animal to a recovery cage, as described above (step 3.1).
   13. Exit Prairie view and ensure all the data is saved/transferred. Shut down the computer and shut down all hardware.
2. Post-processing with FIJI
   NOTE: Fiji is an open-source software focused on biological image analysis²¹.
   1. Download FIJI with respect to the operating system. Unzip the folder contents and run the .exe file.
   2. Drag the .env file gathered from the two-photon imaging to the dialogue box. Split the channels into different image boxes. This will create two different images with different channels, one containing the cell signal and the other the particle signal.
   3. Copy one of the images and click on File > New > Internal Clipboard to create another image. Rename one of the images as Source and the other as Copy.
   4. Use the copy image and click on Process > Enhance contrast in the dialogue box. Click Normalize and OK and then Process > Smooth. This image is used to create a mask and not for analysis. Click on Image > Adjust > Threshold in the dialogue box. Use the sliding scale and method that is appropriate to extract, then click Apply.
   5. Use Process > Binary > Erode to erode single pixels in the background and click Process > Binary > Dilate to add back pixels to the image.
   6. Click on Process > Image Calculator in the dialogue box, select the source image as the first and select the second image as Copy. Use AND to create the intersection of both images. Repeat the same step for the other channel image.
   7. For signal analysis outside of the vascular region, open an unaltered copied image and delete the vascular area of the signal with the Freehand ROI tool.
   8. Set desired parameters by going to Analyze > Set Measurements. Make sure Area, Integrated Density, and Mean Grey Value are checked.
   9. Now analyze by Analyze > Measure. A window with the measurements will pop up. Copy the data into a spreadsheet.
   10. Now select a small area of the image that has no fluorescence. This will be the background.
   11. Click Analyze > Measure for that region. Copy data into a spreadsheet.
   12. Save the resulting image as File > Save as > Analyze file type for remeasurements or publication purposes.

Results

Here, we performed cranial window surgery and engrafted C6 cells in an NSG mouse model of GBM (n = 5). A proper seal between all components involved in the creation of the window (Figure 1A) will ensure the windows' durability for long-term imaging and, additionally, reduce morbidity. Using the stage adapted for in vivo 2P-IVM (Figure 2), we could image animals under anesthesia for up to 2 h without any major motion artifacts. Approximately 10 min a...

Discussion

We present a method for real-time in vivo NP tracking using 2P-IVM through a cranial window to evaluate the tumor delivery of fluorescent-labeled iron oxide NPs. The surgical technique for this procedure requires a steady hand and advanced experimental surgical skills. It is advisable to practice using carcasses or phantoms before moving forward to live animal experience. As an alternative, Hoeferlin et al. implemented a robotic drill to reduce thermal damage, minimize surgical technique variability, and standar...

Materials

Name	Company	Catalog Number	Comments
0.9% sodium chloride infusion solution	Baxter Corp	533-JB1301P
Dulbecco's Modified Eagle Medium	Invitrogen	11965-092
1 mL syringes	BD	Luer-Lok syringe, REF309628
10% FBS	Thermo fisher	Cytiva SH30910.03HI
10% DMSO	Sigma-Aldrich	D8418-50ML
2-photon microscope	Prairie Technologies, Bruker	Prairie Ultima IV
Alcohol applicators, 70%	Medline Industries, LP	MDS093810
Alcohol, spray bottle	Decon Labs Inc	Decon SaniHol, 04-355-122
Aluminum foil	Reynold Brands	Reynold Wrap non-stick aluminum foil
Anesthesia machine	Patterson Scientific	SAS3
Anesthesia monitoring	Kent Scientific	MouseSTAT Jr. Rodent Pulsoxymeter
Antibiotic-Antimycotic (100x), liquid	Invitrogen	15240-096
Betadine applicators	Professional Disposables International, Inc	S41125
Biopsy punch, 5 mm	Miltex	Size 5
Buprenorphine sustained release	Zoo Pharm	Bup SR Lab, 1.0 mg/mL	A generic drug can be used instead.
C6 rat glioma cell line	ATCC (American Tissue Culture Collection)	CCL-107
Cannulas	BD	16 G, 1.1/2”, 30 G, 1”
Carprofen	Pfizer	Rimadyl, 50 mg/ml	A generic drug can be used instead.
Cefazoline	Sagent Pharmaceuticals	25021-101-10, 1 g/vial	A generic drug can be used instead.
Cell strainer, 40 µm	Fisher Scientific	87711
Cotton tip applicators, 6”	Dyad Medical Sourcing, LLC	HCS1005
Dental cement	Stoelting Co	51459	Dental cement kit, clear, 2 components
Dexamethasone	Bimeda	138RX, 2 mg/mL	A generic drug can be used instead.
DietGel	ClearH2O	Recovery, 72-06-502
Drape	Cardinal Health	Bio Shield Wrap
Drill	Saeyang Microtech	Escort Pro, B08350
Drill tips	Hager & Meissinger GmbH	REF310104001001005	Size 005, US 1/4
FIJI imaging analysis software	National Institute of Health	https://imagej.net/software/fiji/
Forceps	Fisher Scientific	13-812-41
Gauze	Fisher HealthCare	Sterile Cotton Gauze Pad, 4 x 4”, 22-415-469
Gelfoam	Ethicon Inc.	Surgifoam absorbable gelatin sponge, Ref. 1972
Germinator	Cellpoint Scientific	Germinator 500, No. 11688
Glass coverslips, 5 mm diameter	Fisher Scientific	Menzel Cover glass
Gloves, non-sterile	Fisher Scientific	Nitrile powder-free medical examination gloves
Gloves, sterile	Medline Industries, LP	MDS104070
Hair removal cream	Church & Dwight	Nair Hair remover lotion
Hamilton syringe	Hamilton Company Inc	Gastight #1701, 10 µL
HBSS without Ca, Mg	Fisher Scientific	PI88284
Head bar	Hongway	5 mm inner diameter O-rings
Heating pad	Stoelting Co.	Rodent warmer X2
Insulin syringes	Exel International Medical Products	29G x 1/2″
Iron oxide nanoparticles	Covis Pharma GmbH	Feraheme ferumoxytol injection, 510 mg/17 mL, 59338077501
Isoflurane	Dechra	26675-46-7
Mice	Jackson Laboratories	NSG, Strain 005557
Microscope (surgery)	Seiler Medical	Seiler IQ Q-100-220
Nanoparticles	Custom		Iron oxide nanoparticles (Ferumoxytol) labeled with fluorescein isothiocyanate
Ophtalmic ointment	Major pharmaceuticals	Lubrifresh P.M. nighttime ointment, 203964
Oxygen	Linde Gas & Equipment Inc.	High Pressure Steel K Style Cylinder, 249CF, 2000PSIG, CGA 540
Plastic cups	Georgia-Pacirif Consumer Products	Dixie Portion Cup, 2 oz., Plastic, Clear, PK2400
Polyethylene tubing	Braintree Scientific	50-195-5494
Scale	Ohaus Corp	CR2200
Scalpel	Integra Life Sciences Production Corp	Integra Miltex Stainless steel disposable scalpel
Scissors	Fisher Scientific	13-804-18
Sealant	Henkel Corp	Loctite 4014
Single use lab gown	High Tech Conversions	17-444-081
Stereotactic frame	Stoelting Co.	Stoelting New Standard TM
Sterile Vacuum Bottle Top Filtration Systems	Fisher Scientific	S2GPU05RE, MilliporeSigma NO.:S2GPU05RE
Styrofoam box	N/A	N/A
Surgical gloves	Cardinal Health	19-163-108
Surgical glue, 3M Vetbond tissues adhesive	3M Animal Care Prodcuts	1469SB
Tail vein cathether	Custom		Consists of two 30 G cannulas connected with sillicone tubing
TrypLE Express (1x), no phenol red	Invitrogen	12604-039
Ultraviolett torch	Spring sunshine technology	Consciot