In Vivo Microdialysis Method to Collect Large Extracellular Proteins from Brain Interstitial Fluid with High-molecular Weight Cut-off Probes

Summary

In vivo microdialysis has enabled collection of molecules present in brain interstitial fluid (ISF) from awake, freely-behaving animals. In order to analyze relatively large molecules in ISF, the current article specifically focuses on the microdialysis protocol using probes with high molecular weight cut off membranes.

Abstract

In vivo microdialysis is a powerful technique to collect ISF from awake, freely-behaving animals based on a dialysis principle. While microdialysis is an established method that measures relatively small molecules including amino acids or neurotransmitters, it has been recently used to also assess dynamics of larger molecules in ISF using probes with high molecular weight cut off membranes. Upon using such probes, microdialysis has to be run in a push-pull mode to avoid pressure accumulated inside of the probes. This article provides step-by-step protocols including stereotaxic surgery and how to set up microdialysis lines to collect proteins from ISF. During microdialysis, drugs can be administered either systemically or by direct infusion into ISF. Reverse microdialysis is a technique to directly infuse compounds into ISF. Inclusion of drugs in the microdialysis perfusion buffer allows them to diffuse into ISF through the probes while simultaneously collecting ISF. By measuring tau protein as an example, the author shows how its levels are altered upon stimulating neuronal activity by reverse microdialysis of picrotoxin. Advantages and limitations of microdialysis are described along with the extended application by combining other in vivo methods.

Introduction

ISF comprises 15-20% of total brain volume and offers a microenvironment critical for signal transduction, substrate transport and waste clearance¹. Therefore, the ability of collecting ISF from living animals will provide greater implications for various biological processes as well as disease mechanism. In vivo microdialysis is one of the few methods that sample and quantify extracellular molecules from ISF from awake, freely moving animals and thereby serves as a useful tool in neuroscience research field^2,3. In this method, microdialysis probes with semipermeable membranes are inserted in the brain and perfused with perfusion buffer at the relatively slow flow rate (0.1-5 µL/min). During this perfusion, extracellular molecules in ISF passively diffuse into the probe according to the concentration gradient and collect as a dialysate. Although this article focuses on the method to sample ISF in the brain, both the principle and the method can be applied to other organs by appropriate modification if necessary.

Microdialysis was first employed in the early 1960s, and since then it has been extensively used to collect small molecules including amino acids or neurotransmitters in brain. However, recent commercial availability of microdialysis probes with high-molecular weight cut off membranes (100 kDa-3 MDa) has extended its application to relatively larger proteins in ISF as well^4,5,6,7. The studies using these probes has led to the finding that proteins such as tau or α-synuclein that were long thought to be exclusive cytoplasmic are also physiologically present in ISF^4,5,8.

One of the difficulties using microdialysis probes with large cut off membranes (typically over 1,000 kDa) is that they are more susceptible to ultrafiltration fluid loss due to the inner pressure accumulated in the probes. Microdialysis probes used here have a unique structure to avoid this issue. The pressure will not be built up due to this structure, thus microdialysis with these probes should be operated in a "push-pull" mode using a syringe pump to perfuse the probes (=push) and a roller/peristaltic pump to collect the dialysate coming from the probe outlet (=pull)⁹ (Although it needs both push and pull pumps, due to pressure cancelling vent holes present in the probes, the system is technically only driven by the pull pump). This article starts with the stereotaxic surgery of a guide cannula implantation and describes how to set up microdialysis lines in order to collect ISF through microdialysis probes with 1,000 kDa cut-off membranes.

Protocol

All animal studies were reviewed and approved by the Institutional Animal Care and Use Committee of the Graduate School of Medicine at the University of Tokyo.

1. Pre-surgical Procedure

1. Before starting surgery, wipe everything with 70% ethanol to maintain sterile conditions. Thermal support using a heating pad is recommended.
2. Anesthetize the mice by intraperitoneal injection of chloral hydrate (400 mg/kg). Confirm anesthetization by performing a toe pinch. Use of meloxicam SR at induction and buprenorphine upon recovery BID for at least 24 h is recommended.
3. Shave the hair with a surgical clipper. Fix a mouse in the mouse and neonatal rat adaptor using ear bars and a nose clamp.
   NOTE: It is critical to make sure that mouse head is secured at this stage and does not move side-by-side.
4. Apply vet ointment on eyes to prevent dryness while under anesthesia.

2. Stereotaxic Surgery for Guide Cannula Implantation

1. Set the mouse and neonatal rat adaptor on the stereotaxic apparatus. Make an incision sagittally on the skin over the skull using a scalpel.　Wipe off blood and the connective tissue on the skull using a damp cotton swab.
2. Determine the coordinate for brain region of interest using brain atlas. Before starting measurements, make sure that midline is straight so that the drill bit can be moved A-P and remain on the midline suture the entire time.
   NOTE: This article uses the coordinate (A/P: -3.1 mm, M/L: -2.5 mm, D/V: -1.2 mm, 12 degrees) to target posterior hippocampus.
3. Level skull A-P
   1. Attach a drill on a manipulator of stereotaxic frame.Lower the drill until it gently touches lambda and record its ventral coordinate. Repeat this procedure for bregma.
      NOTE: When the skull is leveled, the vertical measurement of bregma is equal to that of lambda. If not, adjust the height of nose clamp accordingly. After the skull gets leveled, record the anterior/posterior, lateral coordinate of bregma.
4. Level skull left-right
   1. Move the drill from bregma to the coordinate (A/P: -3.1 mm, M/L: +2.0 mm), lower the drill to the skull and record the vertical coordinate. Then repeat this procedure for the coordinate (A/P: -3.1 mm, M/L: -2.0 mm).
      NOTE: If the skull is leveled, these vertical measurements of two equidistant points from the midline are equal. If not, adjust the height of the ear bars.
5. Drill a burr hole carefully at the target coordinate (A/P: -3.1 mm, M/L: -2.5 mm) to implant a guide cannula. If the diameter of a burr hole is not large enough for a guide cannula implantation, drill another hole that overlaps with the first one. Drill another hole on the right (contralateral side) parietal bone and insert a bone screw, which helps to secure dental cement in 1.10 (See Figure 1B).
6. Cut a circular locking piece from the back side of lid of a 1.5 mL centrifuge tube by a razor blade and make a ''crown''. This crown is used to prevent dental cement from spreading to the skin. Place it on the skull so that the burr holes made in step 2.5 stay within the circle (See Figure 1).
7. Set up a stereotaxic assembly by putting a guide cannula on the shorter arm of a stereotaxic adaptor and fasten it using a cap nut. Set the longer arm of the stereotaxic adaptor on the electrode clamp. Attach it on the manipulator of the stereotaxic apparatus (See Figure 1A).
8. Rotate the D-V stereotax assembly on the manipulator arm by 12 degrees (See Figure 1B). Move the guide cannula to the burr hole made in 1.6. Insert the guide cannula slowly into the brain by lowering it by 1.2 mm.
   NOTE: The angle of the probe is specific to the hippocampus; other regions may require other angles or no angle at all. Consult a brain atlas for precise coordinates. Dry the surface of the skull, because if not the cement will not stick and cement cap can become dislodged
9. Add the dental cement within the crown to fully cover both the metal part of the guide cannula and the bone screw enough to secure them. Apply additional dental cement if there is a part of skull exposed.
10. Wait until dental cement is completely dried (~12-20 min). Remove the stereotaxic adaptor from the electrode clamp. Remove the cap nut and replace the stereotaxic adaptor with a dummy probe and fasten the cap nut.
11. Release the mouse from the stereotaxic apparatus and house the mouse alone in an individual cage.
    NOTE: The mouse should not be left unattended until it has regained sufficient consciousness to maintain sternal recumbency. Check the mouse daily until the day of microdialysis. The mouse receives NSAIDs such as carprofen if it appears to be in pain. Waiting 2 weeks is necessary for sleep-wake studies so the mouse habituates to the new environment¹⁰, but other types of studies may require shorter recovery periods (e.g., 1-2 days).

3. Microdialysis Setup

1. Quality-check of probes: Fill a disposable 1ml syringe with distilled waterand connect it to the outlet (shorter port) of a probe using a byton tube. Cover the vent holes with the fingers and depress the syringe plunger gently to infuse water to the probe. Check that water appears from the probe inlet and there is no leakage on the surface of a microdialysis membrane.
   1. Activation of probes: Submerge the membranes of a probe in ethanol (70-100%) for two seconds. Then, infuse distilled water into the probe again with a syringe again.
2. Preparation of perfusion buffer: In order to avoid adhesion of target molecules to the tubings,add BSA by diluting 30% BSA solution to 4% with artificial CSF (1.3 mM CaCl₂, 1.2 mM MgSO₄, 3 mM KCl, 0.4 mM KH₂PO₄, 25 mM NaHCO₃, 122 mM NaCl, pH=7.35), which closely matches electrolyte concentration in CSF, immediately prior to use. Filter the perfusion buffer through a syringe filter unit with 0.1 µm pore size .
   NOTE: 4% BSA improves recovery for sticking proteins but can severely limit delivery of compounds, especially for compounds that have high BSA-binding. This is one advantage of using lower concentration of BSA (such as 0.15%) in certain instances³. Note that BSA can aggregate very easily when agitated by vortex or stir plate. These aggregates can clog probes or membrane pores. Use caution when preparing a BSA solution to limit this aggregation
3. Prepare two separate lines for inlet and outlet (See Figure 1C) and connect both lines with a connection needle. Fill a disposable 3 mL syringe filled with perfusion buffer and connect it with the inlet end of the tubing using a blunt-end needle. Fill the entire tubing with perfusion buffer by running the syringe pump.
4. Stop the syringe pump and replace the connection needle between inlet line and outlet line with an activated microdialysis probe from step 3.3 (See Figure 1C). Before this replacement, put the cap nut on the probe.
5. Mount the roller tube in the outlet tubing on the roller pump. Start the syringe pump at 10 µL/min and then the roller pump at the slightly slower flow rate (9.5-9.8 µL/min). Make sure to remove all air bubbles in the entire tubing, which may influence the recovery when they enter the probe.
6. Anesthetize the mouse from step 1.12 in the same way as in step 1.2. Put the mouse collar around its neck. Remove the cap nut and dummy probe and slowly insert a microdialysis probe from 3.4 through the guide cannula and fasten the cap nut.
7. Place the mouse in the cage connected to a free-moving system and tether the mouse with the collar. Keep running both syringe pump and roller pump at the indicated flow rate in step 3.5 for at least 1 h.
   NOTE: To achieve ISF collection from awake animals, this article uses a free-moving system, where the cage itself responds to an animal's movement to keep tubings from twisting. Alternatively, liquid swivels available from various companies can be also used.
8. Stop the roller pump first and then the syringe pump. Set the desired flow rate. Run the syringe pump 20% faster than the roller pump.
   NOTE: For instance, if you run at 1 µL/min, operate the syringe pump at 1.2 µL/min. Optimal flow rate should be determined empirically for each molecule.
9. Collecting ISF samples: place the free end of outlet tubing on the refrigerated fraction collector.
   NOTE: Appropriate sample volume varies depending on assays used for analysis.
10. After the completion of the experiment, remove the probe. (Anesthetize the mouse as needed.)  Handle mouse recovery the same way as in step 2.11. Analyze the collected ISF by methods such as HPLC or ELISA.
11. To wash the entire tubing after microdialysis, connect inlet and outlet tubings again by replacing the probe with the connection needle and run the diluted bleach in the entire tubing and then flush it with water. Dry and store it for repeated use.
    NOTE: Other types of tubings are acceptable, however, BSA in perfusion buffer can clog the tubings with small diameter, thus these tubings are considered as single-use. The tubings can be worn down or clogged after multiple uses, so make sure that the flow rate is consistent every time before use.

Results

To stimulate or inhibit neuronal activity in reverse microdialysis^11,12,13, picrotoxin, GABA_A receptor antagonist or tetrodotoxin, Na⁺ channel blocker have been used. It has been shown that tau release is stimulated by increase of neuronal activity^13,14. Consistent with these previous observations, when 50 µM picroto...

Discussion

Microdialysis with high molecular weight cut off membranes has to be operated by a push-pull mode, thus it is critical that the flow rate is accurate and constant. The inaccuracy in the flow rates can be the cause of air bubble generation and inconsistency in the sample concentration. If the flow is inconsistent, check all connections for leakage. If the problem still persists, it may be necessary to re-start with new probes and tubings.

Microdilaysis probes are continuously perfused by the pe...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
The Univentor 820 Microsampler	Univentor	8303002	Refrigerated fraction collector
Syringe pump	KD scientific	KDS-101
Roller pump	Eicom microdialysis	ERP-10
Raturn Stand-Alone System	BASi	MD-1409	Free-moving system
Dual species cage kit	BASi	CX-1600
AtmosLM Microdialysis probe (shaft length 8 mm, membrane length 2 mm)	Eicom microdialysis	PEP-8-02	Shaft length for a probe, a guide, a dumy probe and a stereotaxic adaptor should be identical.
Microdialysis guide (shaft length 8 mm)	Eicom microdialysis	PEG-8
Microdialysis dummy probe (shaft length 8 mm)	Eicom microdialysis	PED-8
Bone screw	BASi	MD-1310
Super bond C&B set	Sunmedical		Dental cement
Small animal Stereotaxic Instrument with digital display console	Kopf	Model 940	Stereotaxic apparatus
Mouse and neonatal rat adaptor	Stoelting	51625
Standard Ear Bars and Rubber Tips for Mouse Stereotaxic	Stoelting	51648
Albumin solution from bovine serum	Sigma	A7284-50ML	30% BSA solution
FEP tubing (70 cm)	Eicom microdialysis	JF-10-70	Internal volume = 0.5 µL/cm
Teflon tubing (50 cm)	Eicom microdialysis	JT-10-50	Internal volume = 0.08 µL/cm
Byton tube	Eicom microdialysis	JB-30
Intramedic luer stab adaptor 23G	BD	427565	Blunt end needle
Roller tube	Eicom microdialysis	RT-5S	Internal volume = 4 µL
Cap nut	Eicom microdialysis	AC-5
0.25 mL microcentrifuge tube with cap	QSP	503-Q	Tubes for fraction collector
Sterotaxic adaptor (shaft length 8 mm)	Eicom microdialysis	PESG-8
Connection needle	Eicom microdialysis	RTJ
Mouse animal collar	BASi	MD-1365
High Speed Rotary Micromotor kit	FOREDOM	K.1070	Drill
Picrotoxin	Sigma	P1675
Screw driver for bone screws
Scalpel
Cotton swab
Surgical clipper