Photobiomodulation Under Electroencephalographic Controls of Sleep for Stimulation of Lymphatic Removal of Toxins from Mouse Brain

All procedures were performed in accordance with the "Guide for the Care and Use of Laboratory Animals", Directive 2010/63/EU on the Protection of Animals Used for Scientific Purposes, and the guidelines from the Ministry of Science and High Education of the Russian Federation (Nº 742 from 13.11.1984), which have been approved by the Bioethics Commission of the Saratov State University (Protocol No. 7, 22.09.2022).

1. Hardware assembly

1. Cut a piece of foil textolite that is 1.5 mm thick to the dimensions of 1.5 mm x 2 mm. This part will be referred to as the light-emitting diode (LED) printed circuit board (PCB) (Figure 1C).
2. Solder an LED to a PCB, as shown in Figure 1.
3. Solder two stranded copper wires to the pins of an LED and then cover them with a sleeve.
4. Print out the 3D model of the frame (Figure 1A); place an LED (Figure 1B) and magnets (Figure 1D) on the frame; place a washer (Figure 1E) on the mouse's head.
5. To assemble the circuit (Figure 2), follow these steps.
   1. First, connect the resistor (Figure 2; R1) between an LED anode and 5 V port on the Arduino.
   2. Next, connect an LED cathode to the metal-oxide-semiconductor field-effect transistor (MOSFET) (Figure 2; Q1) drain. Then, connect the MOSFET source to the ground.
   3. Connect the pull-down resistor (Figure 2; R2) between the MOSFET gate and the ground. Finally, connect the MOSFET gate to pin 3 on the Arduino.
6. Connect a liquid crystal display (LCD) keypad shield to the Arduino.
7. Print out the 3D models of the case, the cover plate, and the buttons. Insert the Arduino board with LCD keypad shield, MOSFET, and LED connector into the case, as shown in Figure 3.

2. Software guide (Figure 4)

1. Download the Arduino sketch (.ino file) and open it via the Arduino integrated development environment (IDE) (Supplementary Coding File 1).
2. Select the correct communication (COM) port and flash the firmware.
3. The interface includes two columns. Use the buttons Left and Right to navigate between these columns. The selection indicator is located to the left of the column.
4. The left-hand column on the screen is the pulse width modulation (PWM) duty cycle selection field. Use the Up and Down buttons to adjust the duty cycle. Achieve the 10/20/30 Dj/cm² dose of PBM with a corresponding 2%/4%/6% PWM duty cycle in a 17 min session.
5. The right-hand column on the screen is the RUN/Off field. To select this column, press the Right button on the keypad and then press Select. When the process is running, the activity indicator will blink, and the text on the RUN button will change to OFF.
   NOTE. Mice are prepared for the experiments over a period of 10 days, including implantation of EEG electrodes, implantation of a chronic catheter into the right lateral ventricle for injection of fluorescent Aβ, and placement of a plate for PBM.

3. Implantation of an EEG recording system (Figure 5)

1. Weigh the mouse and anesthetize it with a mixture of Zoletil 100 and Xylanite (100 mg/kg; 10 mg/kg, respectively) by intramuscular injection into the thigh.
2. When the rear foot withdrawal reflexes and tail pinch response cease, place the mouse in a stereotaxic frame over a heating pad (Figure 5A).
3. Apply ophthalmic ointment to the eyelids to prevent drying of the eyeballs during surgery. Repeat this procedure whenever necessary.
4. Shave the head in the area from the nasal bones to the occipital bones using a shaving machine and sterilize exposed skin first with alcohol.
5. Using straight dissecting scissors, cut off the scalp, hold it with micro forceps, clean the skull from fascia, and dry it with cotton swabs. If necessary, use the hemostatic agents.
6. Using a drill with a diameter of 1.3 mm, make 2 holes in the temporal bones on each side along the coordinates: AP = -1 mm for the first pair of screws and AP = -3 mm for the second pair of screws.
7. Place the EEG screws with wire leads in alcohol for 5 min. Afterward, place the EEG screws in the saline solution.
8. Place four silver-plated screws with electrodes into the holes to a depth of 1 mm (Figure 5B).
9. Fix the screws on the surface of the skull using dental acrylic so that the electrodes extending from them are located towards the animal's nose (Figure 5C). Allow the dental acrylic to harden for 15 min.
10. Attach an EEG recording sensor to the animal's nose using dental acrylic. Allow the dental acrylic to harden for 30 min (Figure 5D).
11. Place the EMG electrodes on the back of the orbicularis oculi muscle using curved tweezers and fix them with the dental acrylic. Allow the dental acrylic to harden for 15 min (Figure 5E).
12. Connect the EEG electrodes to the silver-plated recess of the sensor and solder them using a soldering station. Afterward, fix the EEG electrodes using the dental acrylic (Figure 5F).
13. After surgery, place the mouse on a heating pad to maintain body temperature until the animal fully recovers from anesthesia.
14. Afterward, put the mouse in an individual home cage with free access to food and water with ibuprofen (40 mg/kg in 200 mL of water) for analgesia after surgery for 10 days.

4. Implantation of a plate for PBM

1. Weigh the mouse and anesthetize it with a mixture of Zoletil 100 and Xylanite (100 mg/kg; 10 mg/kg, respectively) by intramuscular injection into the thigh 7 days after implantation of the EEG recording system.
2. When the rear foot withdrawal reflexes and tail pinch response cease, fix the mouse in a stereotactic system.
3. Apply ophthalmic ointment to the eyelids to prevent drying of the eyeballs during surgery. Repeat this procedure whenever necessary.
4. Fix a metal plate with a diameter of 5 mm on the occipital bone of the skull using dental acrylic and Dumont forceps. Allow the dental acrylic to harden for 15 min (Figure 6).

5. Preparation of a chronic catheter

1. Mark on the insulin needle a segment 2 cm from the side of the beveled end.
2. Fix the insulin needle in the needle holder from the side of the beveled tip to the marked segment.
3. Place a 2 cm-PE-10 polyethylene catheter over the entire remaining length of the needle.
4. Fill the catheter with a saline solution and cover it with a plastic cap to seal it.

6. Implantation of a chronic catheter into the right lateral ventricle

1. After implantation of the plate for PBM, make a trepanation hole at the coordinates AP = -0.5 mm and ML = 1.2 mm, with a diameter of 1.5 mm, using a drill.
2. Place the PE-10 polyethylene catheter in a stereotactic holder and insert it into the mouse skull (DV = 2 mm). Afterward, fix it using the dental acrylic. Allow the dental acrylic to harden for 15 min (Figure 7).
3. After surgery, place the mouse on a heating pad to maintain body temperature until the animal fully recovers from anesthesia.
4. Afterward, put the mouse in an individual home cage with free access to food and water with ibuprofen (40 mg/kg in 200 mL of water) for analgesia after surgery for 10 days.

7. tPBM under EEG control of NREM sleep

1. Connect any commercial EEG recording system to the connector on the mouse's head and set the requirement value of the PWM duty cycle.
2. Monitor the EEG signal and wait for delta rhythm activity. If NREM sleep is seen, initiate the PBM process and stop the process if NREM sleep transitions to rapid eye movement (REM) sleep or wakefulness. The dose increases with each interaction until it reaches the required value.
3. When the required PBM dose is obtained, the session is over.

8. Confocal imaging of lymphatic removal of Aβ from mouse brain

1. Connect a 10 cm catheter to an insulin needle.
2. Using a Hamilton syringe with a 29 G needle, prepare an infusion of fluorescent beta-amyloid (FAβ) in a volume of 5 µL into the catheter. Let the catheter remain on a Hamilton syringe.
3. Fix the mouse's hand and connect the catheter through a needle to the implanted chronic catheter.
4. Connect the catheter to a microinjector. Afterward, place the mouse in an individual box;
5. Select the injection rate 0.1 µL/min in the microinjector menu, and press the Start button.
6. Make injection of FAβ into the right lateral ventricle.
7. After FAβ administration, make PBM using an LED for 61 min following the algorithm: 17 min – light and 5 min – pause over 61 min.
8. After PBM, intravenously inject any tracer for labeling the cerebral vessels via the tail.
9. After injection, euthanize mice using the CO₂ euthanasia chamber.
10. Using sharp, straight scissors, make a small transverse incision in the skin along the trachea, holding the skin with straight non-sharp tweezers.
11. Make a longitudinal incision along the entire length of the neck using straight scissors.
12. Using curved tweezers, take up the salivary glands and carefully separate them from the connective tissue. Place a wound retractor on the open section of the incision and fix it in such a way as to push back the surrounding tissues.
13. Examine the area between the trachea and the cleidomastoid muscle using two curved tweezers from both sides of the neck.
14. When the deep cervical lymph node (dcLN) is detected on each side, take off dcLNs using straight tweezers with blunt ends and cut it from the connective tissue.
15. Place dcLNs in a Petri dish with a saline solution and cover them with horizontally oriented cover glass (25 mm × 50 mm × 0.17 mm).
16. Use any commercial confocal microscope to obtain images of whole dcLNs.

9. Analysis of Aβ in the lysates of brain tissues

1. Prepare the samples for the assay.
   1. Euthanize mice using the CO₂ euthanasia chamber.
   2. Decapitate the mouse, remove the skin from its head, and remove the muscles from the skull.
   3. Make two incisions with sharp, straight scissors from the great occipital foramen to the auditory canal.
   4. Using straight tweezers, separate the ventral part of the skull, the occipital bone, and bones forming the middle ear cavities.
   5. Using tweezers, separate the brain from the parietal and frontal bones.
   6. Using straight scissors, remove the upper jaw and cut off the olfactory bulbs.
   7. Place the brain in the physiological solution.
   8. Rinse the brain in cold phosphate-buffered saline to remove excess blood thoroughly and weigh before homogenization.
   9. Prepare a lysing buffer pH 7.2 containing 1.5 mm KH₂PO₄, 8 mm Na₂HPO₄, 3 mm KCl, 137 mm NaCl and 0.1% Tween20, 10 mM EDTA with a freshly prepared protease inhibitory mixture.
   10. Homogenize the brain in fresh lysis buffer (1 mL of lysis buffer for 200-500 mg tissue sample) with a glass homogenizer on ice.
   11. Sonicate a resulting suspension with an ultrasonic cell disruptor till the solution is clarified.
   12. Centrifuge the homogenates at 10,000 × g for 5 min.
   13. Collect the supernatant using a single channel mechanical pipette (100-1000 µL) and assay immediately or aliquot and store at ≤-20 °C.
2. Prepare the following materials: microplate reader with 450 nm ± 10 nm filter; microcentrifuge tubes; single or multi-channel pipettes with high precision and disposable tips; absorbent paper for blotting the microplate; container for wash solution; 0.01 mol/L (or 1x) phosphate buffered saline (PBS); and deionized or distilled water.
3. Prepare the reagents.
   1. Bring the components of the kit and samples to room temperature (RT; 18-25 °C) before use.
   2. Reconstitute the standard with the 1.0 mL of standard diluent, keep for 10 min at RT, and shake gently (not to foam). The standard stock solution is 300 pg/mL. Prepare 5 tubes containing a volume of 0.6 mL of standard diluent and make a triple dilution series.
   3. Set up 5 points (300 pg/mL, 100 pg/mL, 33.33 pg/mL, 11.11 pg/mL, and 3.70 pg/mL) of the diluted standard, and the last tubes with as blank containing only the standard diluent (0 pg/mL).
   4. Quickly spin down the stock solutions of Detection reagent A and Detection reagent B prior to use. Dilute them 100-fold with Assay Diluent A and B to prepare the working concentration.
   5. Dilute 20 mL of the concentrated wash solution (30x) with 580 mL of deionized or distilled water to make 600 mL of wash solution (1x).
   6. Aspirate the needed dosage of the solution with sterilized tips, and do not dump the residual solution into the vial again.
4. Perform the assay.
   1. Determine wells for a diluted standard, blank, and sample.
   2. Prepare 5 wells for standard points and one well for blank.
      1. Add 50 µL each of standard, blank, and sample dilutions into the corresponding wells, respectively. Then, add 50 µL of Detection reagent A to each well immediately.
      2. Shake the plate gently (a microplate shaker is recommended) and cover it with a plate sealer. Incubate the plate at 37 °C for 1 h. Detection reagent A may appear cloudy. Warm the solution to RT and mix gently until it appears uniform.
   3. Aspirate the solution and wash each well with 350 µL of 1x wash solution with the help of a squirt bottle, multi-channel pipette, manifold dispenser, or auto washer. Leave the plate undisturbed for 1-2 min. Snap the plate onto absorbent paper to completely remove the remaining liquid from all wells. Repeat this procedure 3 times.
   4. After the last wash, aspirate or decant any remaining wash buffer. Ensure complete removal of the washing solution by inverting the plate and blotting it against absorbent paper.
   5. Add 100 µL of Detection reagent B working solution to each well and incubate the plate for 30 min at 37 °C after covering it with the plate sealer.
   6. Repeat the aspiration/washing steps for a total of 5 min.
   7. Add 90 µL of substrate solution to each well and cover the plate with a new plate sealer. Incubate for 10-20 min at 37 °C (Do not exceed 30 min) protected from light. After adding the substrate solution, the liquid will turn blue.
   8. Add 50 µL of a stop solution to each well to terminate the reaction. Adding the stop solution will turn the liquid yellow. Tap the plate on its side to mix the liquid. If the color change is inconsistent, gently tap the plate to ensure thorough mixing.
   9. Ensure complete removal of water and fingerprint on the bottom of the plate and no bubble formation on the liquid surface. Then, read the plate in a microplate reader at 450 nm immediately.
5. Calculate the results.
   1. Determine the average of the duplicate readings for each standard, control, and sample. Plot a standard curve with the log of Aβ 1-42 concentration on the y-axis and absorbance on the x-axis.
   2. Draw a best-fit curve through the points, which can be determined by regression analysis.
   3. If diluted samples were used, multiply the concentration obtained from the standard curve by the dilution factor.
      NOTE: For ELISA, a kit for determining Aβ 1-42 was used in this study.