Use of Cerebral Open-Flow Microperfusion for the Longitudinal Collection of Interstitial Fluid in an Animal Model of Glioblastoma

Summary

In vivo, intercellular communications underpinning many central nervous system diseases, such as glioblastoma (GBM), are notoriously difficult to measure and characterize. Here, we describe procedures of cerebral open-flow microperfusion (cOFM) that can be employed to sample interstitial fluid components in a longitudinal animal model of GBM.

Abstract

Diseases of the central nervous system (CNS) manifest through a complex, dynamic network of intercellular communication. Glioblastoma (GBM), the most common and aggressive primary brain tumor, is emblematic of this complexity, with its rapid progression and dismal prognosis measured in months rather than years. GBM's biology is driven by intricate signaling molecule exchanges between neoplastic and stromal cells, underpinning aggressive disease progression. Here, we describe a detailed cerebral open-flow microperfusion (cOFM) protocol in GBM mouse models, enabling real-time longitudinal monitoring of tumor microenvironment dynamics within the interstitial fluid (ISF). Our approach describes the implantation of durable guide cannula head-mounts, their use for intracerebral glioma engraftment directly through the guide, and the collection of high-fidelity cOFM samples for metabolomic and proteomic LC-MS analyses. Crucially, cOFM overcomes the molecular size limitations of traditional microdialysis. Beyond GBM, cOFM methodology promises transformative insights into a spectrum of CNS disorders, including neurodegenerative, epileptic, and neuropsychiatric conditions, through its capacity to provide etiological and treatment-responsive biomarkers within their respective animal models.

Introduction

Investigating central nervous system (CNS) diseases is notoriously challenging due to the complex, multimodal intercellular signaling mediated by the interstitial fluid (ISF)^1,2,3,4. This complexity is particularly pronounced in brain tumors, such as glioblastoma (GBM), where intercellular signals are conveyed between both neoplastic and stromal cells^5,6,7,8,9. In many cases, the aberrant exchange of diverse biomolecules, including metabolites and proteinaceous messengers, contributes to the underlying etiology of CNS pathologies. This investigational challenge is exacerbated, particularly in GBM, by the fact that the molecular composition of ISF is temporally dynamic and evolves throughout disease progression as well as in response to interventional treatments^10,11.

Traditional preclinical GBM investigation methods include in vitro studies and single-endpoint animal models. The nature of these approaches limits their broader clinical applicability due to their inability to capture both the characteristics of the tumor microenvironment and longitudinal temporal changes. Advancements made in the early 1980s overcame these contextual limitations with the development of intracerebral microdialysis (cMD)^12,13,14,15. Since its inception 45 years ago, cMD has established itself as the standard method for longitudinal sampling of interstitial fluid (ISF) within the living brain¹⁶. This is accomplished through the free diffusion and convection of high-concentration ISF solutes through a semipermeable membrane into a continually flowing, low-solute-concentration perfusate, typically artificial cerebrospinal fluid (aCSF)¹⁷. These semipermeable membranes are composed of various biocompatible polymers with common molecular-weight pore size cutoffs ranging from 20 kDa to 3 MDa¹⁸. This minimally invasive procedure enables the longitudinal in situ collection of a diverse array of analytes, ranging from small molecules, such as metabolites or neurotransmitters, up to large proteins, such as antibodies. However, the structural porosity of cMD membranes coupled with the adsorptive nature of its compositional polymers can significantly impact the relative recovery of analytes with particular biochemical or dimensional characteristics, such as lipids or antibodies, respectively^19,20.

Advancements to overcome this limitation of cMD membranes began in the early 2010s with the development of cerebral open-flow microperfusion (cOFM)²¹. Functioning under similar general principles as cMD, cOFM removes biochemical limitations imposed by semipermeable membranes by exchanging them for a plastic-lattice probe with low-adsorption macroscopic openings of approximately 100 µm size²⁰. A multitude of reports have demonstrated the improved utility of cOFM over cMD through the collection of detectable ISF components, including peptide hormones, nanobodies/antibodies, small lipophilic therapeutics, and PEGylated liposomes^{19,22,23,24,25}. Prior reports examining the physiologic tissue reaction to cOFM guide probe implantation show that blood-brain barrier integrity is re-established by 15 days post-implantation, and that no glial scarring is formed up to 30 days post-implantation²⁶. Due to the replacement of microdialysis membranes with macroscopic openings, investigators have also demonstrated the innovative capacity of cOFM guide probes to facilitate the injection of xenografted glioma cells into the brains of immunodeficient rats, such that atraumatic tumor engraftment and growth occur directly around cOFM sampling area²⁷.

Considerations in the use of cOFM in the study of CNS pathologies must still acknowledge its methodological limitations. Owing to its reliance on the passive diffusion and convection of ISF analytes into a flowing perfusate, these limitations revolve around analyte concentration and sample volume¹⁸. Sampled ISF components will be diluted within the collected cOFM perfusate compared to endogenous levels, leading many investigations to compare relative analyte ratios between experimental conditions²⁸. Typical flow rates range from 0.1 µL/min to 1.0 µL/min, equivalent to only 6 µL/h to 60 µL/h. When accounting for tubing dead-space clearance times, the total sampling duration should also be considered. This is particularly important when performing cOFM collections in anesthetized animals, where animal welfare concerns must be carefully addressed. A more detailed description of these and other considerations is available in a previous methods report from the cOFM development team²⁹.

In this manuscript, we provide an updated protocol for employing longitudinal cOFM sampling in the study of neurologic disease, with a focus on GBM. The protocol includes steps for cOFM guide cannula implantation, tumor cell injection, and engraftment through the cOFM guide, as well as two cOFM sampling configurations in awake, freely moving animals. We present representative data demonstrating treatment effects on metabolomic and proteomic ISF components measured by cOFM, the impact of sampling configuration on analyte concentrations, and the efficacy of cOFM-guided tumor cell implantation.

Protocol

All animal procedures within this protocol are approved by the Institutional Animal Care and Use Committee (IACUC) at Northwestern University (protocol ID: IS00021383) and follow National Institutes of Health (NIH) Guidelines. This protocol was developed for use in C57BL/6J mice, both male and female, with a minimum age of 8-10 weeks. The murine glioma cell line used in this protocol, CT2A, was provided by the Seyfried Lab at Boston University and was cultured and prepared following sterile tissue culture conditions.

1. Surgical planning and preparation

1. Animal information: For cOFM experiments, use mice that are a minimum of 8-10 weeks old to allow for complete growth and development. Fixed-position cranial implants may become dislodged or impair proper growth in the still-developing skull and/or brain.
2. Surgical coordinate selection: Use an animal brain atlas to locate the orthogonal coordinates of the region of interest relative to Bregma. For example, glioma cell implantations are typically performed into the striatum to best replicate native GBM microenvironments and maximize the efficacy of subsequent tumor development. The cOFM guide implantation coordinates used in this protocol are mediolateral (ML) = 2.5 mm, anteroposterior (AP) = 0 mm, and dorsoventral (DV) = 3.5 mm.
   NOTE: Stereotactic coordinates can vary between mouse strains and ages. Please use species- and strain-specific atlas coordinates to ensure desired cOFM guide placement.
3. Selection of appropriate cOFM component configurations: Order components for appropriate cOFM configurations in specific total lengths ranging from 2 mm to 22 mm, with open surface lengths of 1 mm to 2 mm.
   ​NOTE: Animals can dislodge improperly implanted cOFM guide head-mounts while performing grooming behaviors. To minimize mechanical leverage upon the head-mount during this activity, we recommend using guides no longer than 2 to 3 mm more than the intended DV coordinate distance.
4. Planning surgical and sampling procedure timepoints for experimental goals: Depending on the disease of interest and the model characteristics, carefully plan out the timing and order of cOFM-related procedures to optimally investigate the underlying biology in question.
   1. If investigating ISF compositional differences between healthy brain and tumor microenvironments, perform cOFM sampling procedures before and after tumor implantation within the same animal.
   2. If investigating effects of clinical interventions on CNS pathologies, perform cOFM sampling procedures before and after administration in parallel animal cohorts receiving either control or experimental treatment.

2. cOFM guide placement surgery

1. Assemble surgical tools for cOFM guide placement surgery ( Table of Materials) and sterilize by autoclave if indicated.
2. Surgical workplace preparation:
   1. Disinfect biosafety cabinet: Perform surgical procedures within a properly disinfected biosafety cabinet. Activate the bio-safety cabinet air flow and clean the interior surfaces with an institutionally approved disinfectant and wipe-down method. Line the bottom surface of the cabinet with sterile drapes.
   2. Prepare stereotactic instruments: Clean the stereotactic frame with a disinfectant and place it in the bio-safety cabinet. Similarly, clean and disinfect the stereotactic-mounted drill and guide holder and place it in the bio-safety cabinet.
   3. Surgical stage heating pad: Use a heating pad on the stereotactic surgical stage to provide supplemental heat while the animal is anesthetized. Clean and disinfect the heating pad surface and place it onto the stereotactic frame where the animal will be situated. Turn on the heating pad to a low heat and place 1-2 sterile drapes on top.
   4. Set out additional surgical instruments and equipment: Aseptically set out additional sterilized equipment into the bio-safety cabinet.
   5. Recovery cage setup: Place a clean recovery cage onto a heating pad set to the lowest possible temperature. Each animal will need to be placed into an individual recovery cage until the animal is fully recovered, thus the total number of recovery cages will depend on surgery performance and recovery times.
3. Presurgical animal preparations (summarized in Figure 1A):
   1. Anesthesia: Administer institutionally approved anesthetic compounds and doses to achieve appropriate anesthetic depth, using the paw pinch reflex test as well as respiratory depth/rate to ensure sufficient anesthesia. For example, intraperitoneal injection of ketamine - xylazine cocktail (100 mg/kg - 10 mg/kg, in saline) is sufficient for achieving appropriate anesthetic depth for C57BL/6J mice.
   2. Analgesia: Administer institutionally approved analgesic compounds to aid in peri- and post-operative pain management. For example, subcutaneous injection of meloxicam (20 mg/kg, in saline) and Buprenorphine SR (1 mg/kg) are appropriate for perioperative pain management.
   3. Hair removal (shaving, depilatory cream): Remove hair from the intended surgical site atop the skull using an electric shaver and/or depilatory cream. If using depilatory cream, ensure it is not left on the skin too long to avoid irritation.
   4. Ophthalmic ointment: Use a sterile cotton-tipped applicator to gently apply ophthalmic ointment to both eyes to prevent corneal drying during surgery.
   5. Animal placement into stereotactic frame: Properly place the animal into the stereotactic frame by inserting incisors into the bite block, inserting ear bars firmly but gently into the ear canals, and fastening the nose clamp over the animal's nasal bone ( Figure 1B). If properly positioned, the animal's skull should be firmly affixed without any movement.
   6. Aseptic surgical site preparations: Wipe down skin around the surgical area with three alternating swabs of alcohol (ethyl- or isopropyl-) and povidone iodine to aseptically disinfect the area for incision.
4. Incision and skull surface preparation:
   ​NOTE: All the following steps should be performed as aseptically as possible. To this end, the primary animal surgeon should wear sterile surgical gloves over standard laboratory gloves before continuing to the incision.
   1. Incision location and length: Use a sterile scalpel to make a sagittal midline incision along the surface of the skull beginning between the eyes and extending caudally 8-10 mm ( Figure 1C).
   2. Skin retraction: Use sterile forceps, cotton-tipped applicators, and/or a scalpel blade to laterally retract skin from the midline incision. Ensure that both Bregma and Lambda are visible and extend the incision if not.
   3. Periosteum incision: Use a sterile scalpel to make an incision into the periosteum and laterally retract the tissue with forceps.
   4. Hydrogen peroxide swab: Clean the skull surface with dilute 1%-3% hydrogen peroxide applied with a sterile cotton-tipped applicator ( Figure 1D). This also aids in the identification of Bregma and Lambda through the sagittal, coronal, and lambdoid cranial sutures.
   5. Self-etching dental adhesive: Use a 0.3 mL to 1.0 mL insulin syringe to evenly apply 20-40 µL of self-etching dental adhesive to the surface of the skull ( Figure 1D). Apply a dental curing light to aid in etching. Once cured, use sterile cotton-tipped applicators and saline to wipe away excess adhesive.
      ​CAUTION: Self-etching dental adhesives, such as Bond Force, are potentially hazardous substances: flammable hazard (GHS02: H225), health hazard (GHS07 and GHS08: H315, H319, H317, H361fd, H336), and environmental hazard (GHS09: H412). Please follow local and institutional guidelines for proper hazardous waste disposal.
5. Locating and drilling burr holes:
   1. Insert a sterilized 0.7 mm drill bit into the microdrill and install the drill into the stereotactic frame with the appropriate holder.
   2. Locate Bregma at the intersection of the sagittal and coronal sutures and use stereotactic micromanipulators to position the drill bit there.
   3. Reposition the drill bit to the cOFM guide coordinate (ML = 2.5, AP = 0.0), engage the drill to 15,000 rpm, and carefully drill a burr hole by adjusting the DV micromanipulator dial (Figure 1E). Make frequent observations of the skull thickness while drilling to prevent damage to the underlying brain. Use physiologic saline during drilling to flush away bone debris and reduce heat between the drill bit and skull.
   4. Repeat drilling procedure for each anchor screw coordinate (Figure 1E). Recommended anchor screw locations are ML = 2.5, AP = -2.5, and ML = -2.5, AP = -2.5.
   5. After burr holes are drilled, detach the microdrill and drill holder from the stereotactic frame.
   6. Use a 30 G needle to gently puncture the dura mater under the cOFM guide burr hole.
6. Install anchor screws using a pair of sterile forceps and an appropriately sized sterile screwdriver (Figure 1F). Depending on the thread pitch of the screw, anchorage is sufficient when the screw is inserted one to two thread lengths deep. Be careful not to drive anchor screws too deeply to avoid damage to the cortex.
7. Install the cOFM guide and cOFM dummy assemblage into the guide holder and mount holder to the stereotactic frame. Position the cOFM guide tip above the burr hole (ML = 2.5 mm, AP = 0.0mm) at a DV height equal to that of Bregma, and slowly descend the cOFM guide through the burr hole to the appropriate pre-determined depth at a rate of 1 mm/min ( Figure 1F). For example, the cOFM guide placement depth with subsequently intended tumor cell implantation is DV = -3.5 mm.
8. Apply cementing agent to the surface of the skull, ensuring it covers the areas under and around both the cOFM guide and anchor screws in thin, uniform layer. Use a dental curing light to harden each layer of cementing agent before applying subsequent layers ( Figure 1G). Continue applying cementing agent until the skull surface between each component is covered, anchor screws are completely covered, and the area under the cOFM guide up to and around the groove is covered. Be careful not to place the cementing agent on or near the locking wedge.
   CAUTION: Cementing agents, such as Tetric EvoFlow, are potentially hazardous substances: health hazard (GHS07: H317, H319, H335, H360). Please follow local and institutional guidelines for proper hazardous waste disposal.
9. Detach the guide holder from the cOFM guide and slowly raise the guide holder with the DV adjustment knob of the micromanipulator. Remove the micromanipulator from the stereotactic frame.
10. Detach the animal from the stereotactic frame by first loosening and removing the ear bars, loosening and retracting the nose clamp, then lifting the animal's incisors from the bite block.
11. Suture skin closed above the dental cementing agent, ensuring that the cOFM guide body remains exposed. Absorbable 4-0 sutures, such as polyglycolic acid (PGA) or polyglactin (Vicryl), are recommended (Figure 1H).
12. Apply triple antibiotic ointment to the skin area adjacent to the original incision.
13. Animal recovery and support:
    1. Transfer the animal to an isolated recovery cage (one animal per cage) and provide supplemental heat from a heating pad set at low.
    2. Optional: Provide the animal with an anesthetic reversal agent to expedite recovery. For example, subcutaneous injections of atipamezole (1 mg/kg, in physiological saline) will expedite anesthetic recovery times from approximately 1 h to approximately 20 min.
    3. Provide continual observation and support until the animal has sufficiently regained consciousness and sternal recumbency. Do not leave animals unattended until they have fully recovered from surgery and anesthesia.
    4. Once the animal has fully recovered, it can be returned to communal housing cages with other animals.
    5. One day after surgery, provide additional analgesic support by administering an institutionally approved analgesic, such as meloxicam (see step 2.3.2).
14. Allow a 14-day recovery time after the cOFM guide implantation surgery to permit complete healing of the blood-brain barrier before performing any additional procedures on the animals.

3. Intracranial tumor cell implantation by cOFM guide

1. Cell culture and preparation:
   1. Culture and expand cells according to cell type-specific protocols. For example, syngeneic mouse glioma cells such as CT2A can be grown in 10 cm dishes or T-75 flasks containing DMEM with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin supplements.
   2. Prepare cultured cells for implantation according to cell type-specific protocols.
      1. For example, with syngeneic CT2A glioma cells, remove growth media, briefly wash with PBS, and incubate cells for 5 min in 0.05% Trypsin (with 0.53 mM EDTA) at 37 °C.
      2. Neutralize trypsin with an equal volume of 1% FBS-containing DMEM, transfer cells to a 15 mL conical tube, and centrifuge at 300 x g for 5 min at 4 °C to pellet cells.
      3. Remove supernatant and resuspend cell pellet in 1 mL of PBS (or physiologic saline). Determine the total number of cells per mL with either a manual or automated hemocytometer. Centrifuge the cell suspension again at 300 x g for 5 min at 4 °C, remove supernatant, and resuspend in an appropriate volume of PBS (or saline) to achieve 50,000 cells per µL. Place cell suspension on ice until ready to implant (step 3.5.2).
2. Assemble surgical tools for intracranial tumor cell implantation surgery (Table of Materials) and sterilize by autoclave if indicated.
3. Surgical workspace preparation
   1. Disinfect biosafety cabinet: See step 2.2.1.
   2. Prepare stereotactic instruments: Clean the stereotactic frame with disinfectants and place it in the bio-safety cabinet. Similarly, clean and disinfect the stereotactic-mounted syringe holder and attach it to the stereotactic frame.
   3. Perform steps 2.2.3.-2.2.5. for the remaining preparatory steps.
4. Presurgical animal preparations:
   1. Perform steps 2.3.1., 2.3.2., 2.3.4., and 2.3.5. for animal preparation.
5. Tumor cell implantation:
   1. Syringe and infusion insert assembly (summarized in Figure 2)
      1. Soak flanged tubing connectors in 70% ethanol for at least 5 min.
      2. Insert the tip of a sterilized 10 µL or 25 µL gastight glass syringe into one end of a flanged tubing connector.
      3. Place the inlet of a cOFM infusion insert into the other end of the tubing connector.
      4. Fix the syringe-infusion insert assembly into the stereotactic-mounted syringe holder.
   2. Load the syringe-infusion insert assembly with an appropriate volume of cell suspension. For example, load 3 µL of cell suspension at 50,000 cells per µL for a total injection of 150,000 cells per animal.
   3. Replacing dummy insert with infusion insert:
      1. Use forceps to remove the locking wedge from the cOFM guide and dummy assembly, then gently lift out the cOFM dummy from the guide.
      2. Use the ML and AP adjustment knobs on the micromanipulator to maneuver the syringe-infusion insert assembly above the cOFM guide. Use the DV adjustment knob to carefully lower the infusion insert into the cOFM guide and secure it in position by reinserting the locking wedge firmly with forceps.
   4. Inject 3 µL of cell suspension from the syringe at a rate of 1 µL/min. Carefully observe the area around the cOFM guide and infusion insert to ensure that no cell suspension effluxes out of the assembly. After the entire volume has been injected, wait an additional 1 min for intracranial pressure to equilibrate.
   5. Replace infusion insert with dummy insert.
      1. Use forceps to remove the locking wedge from the cOFM guide and infusion insert assembly, then gently adjust the DV adjustment knob of the micromanipulator to lift out the cOFM infusion insert from the guide.
      2. Replace the cOFM dummy into the guide and secure it into position by reinserting the locking wedge with forceps.
6. Detach the animal from the stereotactic frame by first loosening and removing the ear bars, loosening and retracting the nose clamp, then lifting the animal's incisors from the bite block.
7. Animal recovery and support:
   1. Transfer the animal to an isolated recovery cage (one animal per cage) and provide supplemental heat from a heating pad set at low.
   2. Optional: Provide the animal with an anesthetic reversal agent (such as atipamezole) to expedite recovery duration, see step 2.13.2.
   3. Provide continual observation and support until the animal has sufficiently regained consciousness and sternal recumbency. Do not leave animals unattended until they have fully recovered from surgery and anesthesia.
   4. Once the animal has fully recovered, it can be returned to communal housing cages with the company of other animals.
   5. On the day after surgery, provide additional analgesic support by administering an institutionally approved analgesic, such as meloxicam (see step 2.3.2).
      ​NOTE: If using luciferase-positive cell lines, bioluminescent imaging (BLI) can be performed 1-2 weeks after cell implantation to verify tumor engraftment success. While physical characteristics such as animal fur or head-mount presence can impede detected luminescence, it can still be used to make rough approximations of tumor presence or absence. Alternatively, MRI can be performed if the anchor screws used in the cOFM guide placement are non-ferromagnetic and MRI compatible.

4. cOFM sampling of cerebral interstitial fluid

NOTE: Given its reliance on passive analyte diffusion, cOFM samples contain diluted analyte concentrations compared to the endogenous ISF environment. Due to this, several methods have been previously proposed to calibrate sample measurements to better reflect in situ concentrations¹⁸. This section describes two different sampling modalities. Step 4.1. describes the configuration mode recommended by the manufacturer (from here referred to as fractionated) that offers greater temporal sensitivity, but at the expense of analyte concentration. Step 4.2. details an alternative mode of configuration we have developed (from here referred to as recirculated) that can achieve greater analyte concentration at the expense of decreased temporal resolution.

1. Fractionated cOFM sampling (summarized in Figure 3A):
   1. Soak flanged tubing connectors in 70% ethanol for at least 5 min to facilitate attachment onto tubing and sampling inserts.
   2. Load perfusate bag with sufficient aCSF perfusate. For example, 680 µL is minimally sufficient for both a 20 min line flush at 10 µL/min and an 8 h sampling at 1 µL/min. However, a minimum volume of 2 mL is recommended to prevent perfusate bag collapse and malfunction.
   3. Prefill both push and pull tubing lines with an appropriate volume of aCSF using gastight glass syringes. For example, Universal Push-Pull Low-Bind Tubing has an inner diameter of 0.25 mm and individual tubing lengths of 100 cm. This equates to approximately 1 µL per 2 cm, and thus approximately 55 µL in the push tubing and 100 µL in the pull tubing.
   4. Correctly install the peristaltic section of the tubing into the pump head of the microperfusion pump as shown in Figure 3A.
      ​NOTE: See BASi instruction manual for additional instructions if needed.
   5. Connect the perfusate bag to the push section of the tubing using the Luer lock connector (Figure 3A, component i).
   6. Connect the sampling insert to the tubing lines using flanged tubing connectors. The sampling insert inlet (longer shaft length) should be attached to the output end of the push tubing, opposite the perfusate bag tube line (Figure 3A, top left). The sampling insert outlet (shorter shaft) should be attached to the input end of the pull tubing, parallel to the perfusate bag tube line (Figure 3A, top right). Place the connected sampling insert into a sterile microcentrifuge tube containing 750 µL of aCSF in preparation for system flush.
   7. Connect the fraction collector needle with a flanged tubing connector to the pull tube output line (opposite end from the sampling insert outlet).
   8. Perform a tubing system flush for 20 min at 10 µL/min to remove any trapped gas bubbles within the tubing system.
   9. Fraction collector setup:
      1. Label and weigh fraction collection tubes, annotating each weight for subsequent comparison and validation of sample volume collected. Place tubes into the corresponding slots of the fraction collector carousel.
      2. Turn on the fraction collector and configure the settings to the desired values. Example settings include # of samples, sample time (m), delay (hours), refrig cooling, and _cannula mode.
         ​NOTE: The discarding of dead-space fluid volume in the pull tube that never contacted orthotopic ISF is approximated by i) tubing length and volume (approximately 2 cm/ µL for OFM-PP2-100-LB), ii) flow rate, and iii) delay (hours). These parameters should be carefully considered when setting up the collection system to ensure proper fraction collection.
2. Recirculated cOFM sampling (summarized in Figure 3B):
   1. Soak flanged tubing connectors in 70% ethanol to facilitate attachment onto tubing and sampling inserts.
   2. Prefill pull tubing line with an appropriate volume of aCSF using gastight glass syringes.
   3. Correctly install the peristaltic section of the tubing into the pump head of the microperfusion pump.
   4. Connect the sampling insert to the tubing lines using flanged tubing connectors. The sampling insert inlet (longer shaft length) should be attached to the output end of the pull tubing. The sampling insert outlet (shorter shaft) should be attached to the input end of the pull tubing. Place the connected sampling insert into a sterile microcentrifuge tube containing 750 µL of aCSF in preparation for system flush.
   5. Perform a tubing system flush for 20 min at 10 µL/min to remove any trapped gas bubbles within the tubing system.
3. Rotating cage system setup:
   ​NOTE: Rotating animal cage systems, such as the Raturn System, facilitate awake animal cOFM sampling by preventing hyper-coiling and tangling of the push and pull tubing. This can also aid in the automation of cOFM sampling by eliminating the need for manually detangling tubes during awake cOFM collection sessions.
   1. Place clean cage bedding evenly across the rotating cage floor and equip the cage with easily accessible food and water sources.
   2. Turn on the cage rotation controller and test its activity by rotating the suspended tether. The cage should counter-rotate relative to the rotational direction imparted upon the tether. Set the rotation controller to standby until the animal is attached to the tether.
4. Connecting animal to cOFM system:
   ​NOTE: Connection and detachment of animals to the cOFM system is most easily performed while the animal is under temporary isoflurane anesthesia. If performed correctly, this process takes no more than 5 min per animal; however, the use of ophthalmic ointment is highly recommended.
   1. Animal anesthesia: Place the animal into the isoflurane anesthesia induction chamber. Begin anesthesia with oxygen flow at 2 L/min and isoflurane set to 3%. Observe the animal until it reaches an appropriate level of anesthesia (immobile with a respiration rate of approximately 1 breath/s). Transfer the animal from the induction chamber to the anesthesia mask set at 2 L/min oxygen flow and 3% isoflurane.
   2. Animal jacketing: Place the animal in an appropriately sized jacket, drawing the forelimbs completely through the arm holes and fastening the jacket securely along the animal's back (shown in Figure 3C).
   3. Replacing dummy insert with sampling insert: Use a pair of forceps to remove the locking wedge from the guide and dummy assembly and carefully lift the dummy out of the guide. Gently install the sampling, insert it into the guide, then reinsert the locking wedge.
   4. Remove the animal from anesthesia and place it into the rotating cage system. Attach the cage tether to the animal jacket and activate the rotating cage system.
5. Performing cOFM sampling:
   1. Set microperfusion pumps to 5 µL/min and perform a 2 min flush to remove any microbubbles that may have resulted from the cOFM sampling insert installation.
   2. Change the microperfusion pump settings to the correctly desired flow rate for pump mode (for example, 1 µL/min).
   3. Activate pump mode. If performing fractionated cOFM sampling, activate the fraction collector.
   4. During the initial stages of cOFM sampling, make close observations of the push and pull tubing to ensure the absence of gas bubbles, as these will impair proper machine function and sample collection and potentially endanger the animal. If gas bubbles are observed, stop system components, detach the animal from cOFM system, and remove gas bubbles by aCSF flush through the tubing.
   5. Throughout the duration of the cOFM sampling process, make additional frequent observations of the cOFM tubing and animal behavior. If animal behavior indicates sudden health changes, immediately stop system components and address the animal health concern. Consult with institutional veterinary staff if necessary.
6. Detaching an animal from the cOFM system:
   1. Deactivate the rotating cage system and detach the cage tether from the animal jacket.
   2. Transfer the animal to the isoflurane anesthesia induction chamber, with sampling insert and cOFM tubing still attached. Begin anesthesia with oxygen flow at 2 L/min and isoflurane set to 3%. Observe the animal until it reaches an appropriate level of anesthesia (immobile with a respiration rate of approximately 1 breath/s). Transfer the animal from induction chamber to the anesthesia mask set at 2 L/min oxygen flow and 3% isoflurane.
   3. Replace sampling insert with dummy insert: Use a pair of forceps to remove the locking wedge from the guide and sampling insert assembly, and carefully lift the sampling insert out of the guide. Gently install the dummy insert back into the guide, then reinsert the locking wedge.
   4. De-jacketing: Detach fasteners from the back side of the animal jacket and gently remove the jacket from the animal.
   5. Animal recovery and observation: Remove animal from isoflurane anesthesia and place into a clean recovery cage, providing continual observation until the animal fully regains consciousness and sternal recumbency. Re-house into a home cage once the animal is fully recovered.
7. Sample collection and storage:
   1. Fractionated sampling:
      1. Remove fraction collection tubes from the fraction collector and place on ice for short-term transport.
      2. Weigh each collection tube and compare to the pre-collection weight to verify the sample volume.
      3. Proceed to downstream analyses of cOFM samples or place them into -80 °C freezer for long-term storage.
   2. Recirculated sampling:
      1. Leaving the pull tube attached to the cOFM sampling insert outlet (shorter shaft), detach from the rest of the setup and place the tube and sampling insert into a new, clean microcentrifuge tube.
      2. Attach a sterile 18G needle to a 1 mL to 5 mL sterile syringe and fill with air.
      3. Carefully insert the needle into the flanged tubing connector at the input end of the pull tube.
      4. Ensuring that the cOFM sampling insert remains in the microcentrifuge tube, press down on the syringe plunger to eject the recirculated cOFM sample volume from the pull tube with air. Make sure that the entire cOFM sample is ejected from the pull tube and into the microcentrifuge tube, detaching the syringe and refilling with air if necessary.
      5. Place the recirculated cOFM sample on ice for short-term transport.
      6. Proceed to downstream analyses or place them into -80 °C freezer for long-term storage.
8. Storing cOFM supplies if repeating cOFM sampling in the same animal:
   ​NOTE: Re-use of cOFM disposables, such as sampling inserts and inlet-/outlet-tubing, is not recommended by the manufacturer.
   1. Flush the cOFM push-pull tubing with approximately 500 µL of sterile aCSF or saline at a rate of no more than 10 µL/min. Follow this with a similar flush of 1-2 mL of 70% ethanol. Finally, use a 5 or 10 mL syringe equipped with an 18G needle and flanged tubing connector to flush the tubing out with air.
   2. Flush inlet and outlet tubes of the cOFM sampling insert with 50 µL of sterile aCSF or saline using a gastight glass syringe equipped with a flanged tubing connector. Follow this with 50 µL of 70% ethanol. Detach the cOFM sampling insert from the tubing connector and store it in a vial filled with 70% ethanol until ready for reuse.

Results

Various evaluative measures to determine experimental success can occur at intermediate or culminating experimental stages, depending on the specific nature of the underlying pathology and investigational animal model. For examinations into GBM biology, bioluminescent imaging (BLI) can be used as an intermediate evaluative measure for the efficacy of cOFM guide-mediated tumor cell engraftment (Figure 4A). However, this is used for simple dichotomous present or absent tumor evaluations, given...

Discussion

The cOFM procedures described in this protocol hold the potential to revolutionize the study of CNS diseases through their ability to longitudinally sample endogenous components of the ISF of pathologic tissue. However, the investigational success of employing this methodology relies upon the proper performance of several critical steps. Of these, the correct and secure placement of the cOFM guide cannula is one of the most important. Improper surgical procedures or attachment strategies can result in the dislodgement of...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.7 mm Drill bit	Fine Science Tools	19008-07	Burr hole drilling
1 to 5 mL syringe	BD	#309646	Recirculated perfusate sample retreival
18 G needle	BD	#305195	Recirculated perfusate sample retreival
30G needle	BD Ultra-Fine	328438	Puncturing dura
4-0 vicryl sutures	Dynarex	9130	Closing surgical would
Alcohol wipes	FisherBrand	22-363-750	Surgical site skin disinfection
Anchor screws	BASi	MD-1310	cOFM head-mount placement
Artificial cerebral spinal fluid (aCSF)	BASi (Joanneum)	MD-2400	Sampling ISF
cOFM dummy insert	BASi (Joanneum)	cOFM-D-#	cOFM head-mount placement
cOFM guide	BASi (Joanneum)	cOFM-GD-#-#	cOFM head-mount placement
cOFM guide holder	BASi (Joanneum)	OFM-STM-1	cOFM head-mount placement
cOFM infusion insert	BASi (Joanneum)	cOFM-I-#	Cell injection
cOFM locking wedge	BASi (Joanneum)	cOFM-LOCK	cOFM head-mount placement
cOFM sampling insert	BASi (Joanneum)	cOFM-S-X	Sampling ISF
Cotton-tipped applicators	FisherBrane	22363160	Fluid absorption, applying hydrogen peroxide
CT2A mouse glioma cell line	Seyfried Lab - Boston University	NA	Engrafted tumor cells
Dental cementing agent	IvoClar	595953US	cOFM head-mount placement
Dental curing light	Henry Schein	5700253	cOFM head-mount placement
Depilatory cream	Veet	3116875	Surgical site hair removal
Electric shaver	Wahl Clipper Corp.	8841	Surgical site hair removal
Fine tipped forceps	Fine Science Tools	5SF	Incision, anchor screw placement
Flanged tubing connectors	BASi (Joanneum)	MD-1510	Cell injection
Fraction collection vials	BASi (Joanneum)	MF-5281	Fractionated sample storage
Gastight glass syringe (25 μL)	Hamilton	80465	Cell injection
Hydrogen peroxide (1-3%)	Walgreens	NDC: 0363-0268-32	Skull surface cleaning
Insulin syringes (0.3 to 1.0 mL) with 31G needles	BD Ultra-Fine	328438	Anesthetic and analgesic administration
Microcentrifuge tubes	Basix	02-682-002	Recirculated perfusate sample retreival
Microdrill and stereotactic holder	Harvard Apparatus	75-1874	Burr hole drilling
Mouse jacket	Lomir	MJ 02	Animal connection to rotating cage system
OFM low-bind tubing	BASi (Joanneum)	OFM-PP2-100-LB	Sampling ISF
OFM-Pump - Microperfusion pump	BASi (Joanneum)	MPP102-PC	Sampling ISF
Ophthalmic Ointment	Dechra	211-38	Peri-sugrical eye lubrication
Passivated metal sampling needle	BASi (Joanneum)	MW-2310	Fractionated sample storage
Perfusate bag	BASi (Joanneum)	OFM-BAG	Sampling ISF
Physiologic saline solution	ICU Medical	0990-7983-03	Skull surface cleaning, aid in burr hole drilling
Povidone-iodine swabs	PDI Healthcare	SKU B40600	Surgical site skin disinfection
Probe holder	Stoelting	51633
Refrigerated fraction collector	BASi (Joanneum)	MD-1201	Fractionated sample storage
Rotating cage system	BASi (Joanneum)	AMD-R Raturn	Automated housing for awake cOFM sampling
Scalpel blades and handles	Fisher Scientific	22079693	Incision
Screw driver			cOFM head-mount placement
Self-etching dental adhesive	Tokuyama		Skull surface priming for cementing agent
Silicone push cap for collection vials	BASi (Joanneum)	MF-5283	Fractionated sample storage
Sterotactic frame	Stoelting	51725