This research was supported by a grant from the Russian Science Foundation (No. 23-75-30001).

All procedures were performed in accordance with the &#34;Guide for the Care and Use of Laboratory Animals&#34;, 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 (N&#186; 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
<ol>
	<li>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).</li>
	<li>Solder an LED to a PCB, as shown in Figure 1.</li>
	<li>Solder two stranded copper wires to the pins of an LED and then cover them with a sleeve.</li>
	<li>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&#39;s head.</li>
	<li>To assemble the circuit (Figure 2), follow these steps.
	<ol>
		<li>First, connect the resistor (Figure 2; R1) between an LED anode and 5 V port on the Arduino.</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Connect a liquid crystal display (LCD) keypad shield to the Arduino.</li>
	<li>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.</li>
</ol>
2. Software guide (Figure 4)
<ol>
	<li>Download the Arduino sketch (.ino file) and open it via the Arduino integrated development environment (IDE) (Supplementary Coding File 1).</li>
	<li>Select the correct communication (COM) port and flash the firmware.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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 A&#946;, and placement of a plate for PBM.</li>
</ol>
3. Implantation of an EEG recording system (Figure 5)
<ol>
	<li>Weigh the mouse and anesthetize it with a mixture of Ketamine and Xylazine (100 mg/kg; 10 mg/kg, respectively) by intramuscular injection into the thigh. Administer Xylazine as pre-operative analgesia at a concentration of 20 mg/mL at a dose of 1.5 mL/kg.</li>
	<li>When the rear foot withdrawal reflexes and tail pinch response cease, place the mouse in a stereotaxic frame over a heating pad (Figure 5A).</li>
	<li>Apply ophthalmic ointment to the eyelids to prevent drying of the eyeballs during surgery. Repeat this procedure whenever necessary.</li>
	<li>Shave the head in the area from the nasal bones to the occipital bones using a shaving machine and disinfect the exposed skin with alternating rounds of chlorhexidine and alcohol 3 times each.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Place the EEG screws with wire leads in alcohol for 15 min. Afterward, place the EEG screws in the saline solution.</li>
	<li>Place four silver-plated screws with electrodes into the holes to a depth of 1 mm (Figure 5B).</li>
	<li>Fix the screws on the surface of the skull using dental acrylic so that the electrodes extending from them are located towards the animal&#39;s nose (Figure 5C). Allow the dental acrylic to harden for 15 min.</li>
	<li>Attach an EEG recording sensor to the animal&#39;s nose using dental acrylic. Allow the dental acrylic to harden for 30 min (Figure 5D).</li>
	<li>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).</li>
	<li>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).</li>
	<li>After surgery, place the mouse on a heating pad to maintain body temperature until the animal fully recovers from anesthesia.</li>
	<li>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. 
	NOTE: Each animal was kept in an individual cage so that the mice could not deform the EEG registration system. Ibuprofen was supplied with water to prevent stress in animals.</li>
</ol>
4. Implantation of a plate for PBM
<ol>
	<li>Weigh the mouse and anesthetize it with a mixture of Ketamine and Xylazine (100 mg/kg; 10 mg/kg, respectively) by intramuscular injection into the thigh 7 days after implantation of the EEG recording system. Administer Xylazine as pre-operative analgesia at a concentration of 20 mg/mL at a dose of 1.5 mL/kg.</li>
	<li>When the rear foot withdrawal reflexes and tail pinch response cease, fix the mouse in a stereotactic system.</li>
	<li>Apply ophthalmic ointment to the eyelids to prevent drying of the eyeballs during surgery. Repeat this procedure whenever necessary.</li>
	<li>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).</li>
</ol>
5. Preparation of a chronic catheter
<ol>
	<li>Mark on the insulin needle a segment 2 cm from the side of the beveled end.</li>
	<li>Fix the insulin needle in the needle holder from the side of the beveled tip to the marked segment.</li>
	<li>Place a 2 cm-PE-10 polyethylene catheter over the entire remaining length of the needle.</li>
	<li>Fill the catheter with a saline solution and cover it with a plastic cap to seal it.</li>
</ol>
6. Implantation of a chronic catheter into the right lateral ventricle
<ol>
	<li>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.</li>
	<li>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).</li>
	<li>After surgery, place the mouse on a heating pad to maintain body temperature until the animal fully recovers from anesthesia.</li>
	<li>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. 
	NOTE: Each animal was kept in an individual cage so that the mice could not deform the EEG registration system. Ibuprofen was supplied with water to prevent stress in animals. Apply antibacterial ointment to the exposed areas of the skull and soft tissues during the rehabilitation period (7 days) to prevent them from drying out.</li>
</ol>
7. tPBM under EEG control of NREM sleep
<ol>
	<li>Connect any commercial EEG recording system to the connector on the mouse&#39;s head and set the requirement value of the PWM duty cycle.</li>
	<li>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.</li>
	<li>When the required PBM dose is obtained, the session is over.</li>
</ol>
8. Confocal imaging of lymphatic removal of A&#946; from mouse brain
<ol>
	<li>Connect a 10 cm catheter to an insulin needle.</li>
	<li>Using a Hamilton syringe with a 29 G needle, prepare an infusion of fluorescent beta-amyloid (FA&#946;) in a volume of 5 &#181;L into the catheter. Let the catheter remain on a Hamilton syringe.</li>
	<li>Fix the mouse&#39;s hand and connect the catheter through a needle to the implanted chronic catheter.</li>
	<li>Connect the catheter to a microinjector. Afterward, place the mouse in an individual box;</li>
	<li>Select the injection rate 0.1 &#181;L/min in the microinjector menu, and press the Start button.</li>
	<li>Make injection of FA&#946; into the right lateral ventricle.</li>
	<li>After FA&#946; administration, make PBM using an LED for 61 min following the algorithm: 17 min - light and 5 min - pause over 61 min.</li>
	<li>After PBM, intravenously inject any tracer for labeling the cerebral vessels via the tail.</li>
	<li>After injection, euthanize mice using the CO2 euthanasia chamber.</li>
	<li>Using sharp, straight scissors, make a small transverse incision in the skin along the trachea, holding the skin with straight non-sharp tweezers.</li>
	<li>Make a longitudinal incision along the entire length of the neck using straight scissors.</li>
	<li>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.</li>
	<li>Examine the area between the trachea and the cleidomastoid muscle using two curved tweezers from both sides of the neck.</li>
	<li>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.</li>
	<li>Place dcLNs in a Petri dish with a saline solution and cover them with horizontally oriented cover glass (25 mm &#215; 50 mm &#215; 0.17 mm).</li>
	<li>Use any commercial confocal microscope to obtain images of whole dcLNs.</li>
</ol>
9. Analysis of A&#946; in the lysates of brain tissues
<ol>
	<li>Prepare the samples for the assay.
	<ol>
		<li>Euthanize mice using the CO2 euthanasia chamber.</li>
		<li>Decapitate the mouse, remove the skin from its head, and remove the muscles from the skull.</li>
		<li>Make two incisions with sharp, straight scissors from the great occipital foramen to the auditory canal.</li>
		<li>Using straight tweezers, separate the ventral part of the skull, the occipital bone, and bones forming the middle ear cavities.</li>
		<li>Using tweezers, separate the brain from the parietal and frontal bones.</li>
		<li>Using straight scissors, remove the upper jaw and cut off the olfactory bulbs.</li>
		<li>Place the brain in the physiological solution.</li>
		<li>Rinse the brain in cold phosphate-buffered saline to remove excess blood thoroughly and weigh before homogenization.</li>
		<li>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.</li>
		<li>Homogenize the brain in fresh lysis buffer (1 mL of lysis buffer for 200-500 mg tissue sample) with a glass homogenizer on ice.</li>
		<li>Sonicate a resulting suspension with an ultrasonic cell disruptor till the solution is clarified.</li>
		<li>Centrifuge the homogenates at 10,000 &#215; g for 5 min.</li>
		<li>Collect the supernatant using a single channel mechanical pipette (100-1000 &#181;L) and assay immediately or aliquot and store at &#8804;-20 &#176;C.</li>
	</ol>
	</li>
	<li>Prepare the following materials: microplate reader with 450 nm &#177; 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.</li>
	<li>Prepare the reagents.
	<ol>
		<li>Bring the components of the kit and samples to room temperature (RT; 18-25 &#176;C) before use.</li>
		<li>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.</li>
		<li>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).</li>
		<li>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.</li>
		<li>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).</li>
		<li>Aspirate the needed dosage of the solution with sterilized tips, and do not dump the residual solution into the vial again.</li>
	</ol>
	</li>
	<li>Perform the assay.
	<ol>
		<li>Determine wells for a diluted standard, blank, and sample.</li>
		<li>Prepare 5 wells for standard points and one well for blank.
		<ol>
			<li>Add 50 &#181;L each of standard, blank, and sample dilutions into the corresponding wells, respectively. Then, add 50 &#181;L of Detection reagent A to each well immediately.</li>
			<li>Shake the plate gently (a microplate shaker is recommended) and cover it with a plate sealer. Incubate the plate at 37 &#176;C for 1 h. Detection reagent A may appear cloudy. Warm the solution to RT and mix gently until it appears uniform.</li>
		</ol>
		</li>
		<li>Aspirate the solution and wash each well with 350 &#181;L 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.</li>
		<li>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.</li>
		<li>Add 100 &#181;L of Detection reagent B working solution to each well and incubate the plate for 30 min at 37 &#176;C after covering it with the plate sealer.</li>
		<li>Repeat the aspiration/washing steps for a total of 5 min.</li>
		<li>Add 90 &#181;L of substrate solution to each well and cover the plate with a new plate sealer. Incubate for 10-20 min at 37 &#176;C (Do not exceed 30 min) protected from light. After adding the substrate solution, the liquid will turn blue.</li>
		<li>Add 50 &#181;L 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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Calculate the results.
	<ol>
		<li>Determine the average of the duplicate readings for each standard, control, and sample. Plot a standard curve with the log of A&#946; 1-42 concentration on the y-axis and absorbance on the x-axis.</li>
		<li>Draw a best-fit curve through the points, which can be determined by regression analysis.</li>
		<li>If diluted samples were used, multiply the concentration obtained from the standard curve by the dilution factor. 
		NOTE: For ELISA, a kit for determining A&#946; 1-42 was used in this study.</li>
	</ol>
	</li>
</ol>

Enhancing Brain Drainage During Sleep with Transcranial Photobiomodulation

The authors have nothing to disclose.

MLVs are an important target for the development of innovative technologies for modulation of the brain&#39;s drainage and removal of cellular debris and wastes from the brain, especially in aged subjects whose MLV function declines1,22. In a homeostatic state, deep sleep is associated with the natural activation of brain tissue cleansing13,14. Therefore, it is obvious to expect that stimulation of MLVs d...

Meningeal lymphatic vessels (MLVs) play an important role in the removal of toxins and metabolites from brain tissues1,2,3. Damage of MLVs in various brain diseases, including tumors, traumatic brain injuries, hemorrhages, and neurodegenerative processes, is accompanied by a decrease in the MLV functions leading to the progression of these pathologies1,2,3,4,5,6. Therefore, the development of methods for the stimulation of MLVs opens new horizons in the emergence of effective technologies for the treatment of brain diseases. Recently, non-invasive technology for effective transcranial photobiomodulation (tPBM) has been proposed to stimulate MLVs and remove toxins such as blood and A&#946; from the brain5,7,8,9,10,11,12. It is interesting to note that deep sleep is a natural factor for the activation of lymphatic drainage processes in the brain13,14. Based on this fact, it is logical to assume that the tPBM of MLVs during sleep may have more effective therapeutic effects than during wakefulness9,11,12,15. However, there are currently no commercial technologies for tPBM during sleep16. In addition, animal experiments to study the therapeutic effects of tPBM are performed under anesthesia, which is required to accurately deliver light to the brain. However, anesthesia significantly affects the brain&#39;s drainage, which reduces the quality of research results17.
A&#946; is a metabolic product of normal neural activity18. As it was established in cultured rat cortical neurons, A&#946; is released from them at high rates into the extracellular space (2-4 molecules/neuron/s for A&#946;)19. There is evidence that the dissolved form of A&#946;, located in the extracellular and perivascular spaces, is most toxic to neurons and synapses20. The soluble A&#946; is rapidly cleared from the human brain during 1-2.5 h21. MLVs are the tunnels for removal of the soluble A&#946; from the brain1,7 that declines with age, leading to the accumulation of A&#946; in the aged brain1,22. There is evidence that extracellular abnormalities of A&#946; levels in the brain correlate with cognitive performance in aging and are associated with the development of Alzheimer&#39;s disease (AD)23,24. Therefore, aged and old rodents are considered alternatives to transgenic models for the study of amyloidosis, including AD25,26.
This study presents an original and portable tPBM technology under electroencephalographic (EEG) control of deep or non-rapid eye movement (NREM) sleep in non-anesthetized male C57BL/6 mice of different ages to stimulate lymphatic clearance of A&#946; from the brain into the peripheral lymphatic system (the deep cervical lymph nodes, dcLNs).

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.1% Tween20</td>
			<td>Helicon,&#160; Russia</td>
			<td>SB-G2009-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Catheter</td>
			<td>Scientific Commodities Inc., USA</td>
			<td>PE-10, 0.28 mm ID &#215; 0.61 mm OD</td>
			<td></td>
		</tr>
		<tr>
			<td>CO2 chamber</td>
			<td>Binder, Germany</td>
			<td>CB-S 170</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal microscop</td>
			<td>Nikon, Japan</td>
			<td>A1R MP</td>
			<td></td>
		</tr>
		<tr>
			<td>Dental acrylic</td>
			<td>Zermack, Poland-Russia</td>
			<td>Villacryl S, V130V4Z05</td>
			<td></td>
		</tr>
		<tr>
			<td>Drill</td>
			<td>Foredom, Russia</td>
			<td>SR W-0016</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont forceps</td>
			<td>Stoelting, USA</td>
			<td>52100-07</td>
			<td></td>
		</tr>
		<tr>
			<td>Evans Blue dye</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>206334</td>
			<td></td>
		</tr>
		<tr>
			<td>Hamilton</td>
			<td>Hamilton Bonaduz AG, Switzerland</td>
			<td>29 G needle</td>
			<td></td>
		</tr>
		<tr>
			<td>Ibuprofen</td>
			<td>Sintez OJSC, Russia</td>
			<td>N/A</td>
			<td>&#160;Analgesic drug</td>
		</tr>
		<tr>
			<td>Insulin needle</td>
			<td>INSUPEN, Italy</td>
			<td>31 G, 0.25 mm x 6 mm</td>
			<td></td>
		</tr>
		<tr>
			<td>Levomekol antibacterial ointment</td>
			<td>Nizhpharm</td>
			<td>D06C </td>
			<td>For external use at a dose of 40 mg/g, 1 time per day</td>
		</tr>
		<tr>
			<td>Micro forceps</td>
			<td>Stoelting, USA</td>
			<td>52102-02P</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcentrifuge</td>
			<td>Gyrozen, South Korea</td>
			<td>GZ-1312</td>
			<td></td>
		</tr>
		<tr>
			<td>Microinjector</td>
			<td>Stoelting, USA</td>
			<td>53311</td>
			<td></td>
		</tr>
		<tr>
			<td>Non-sharp tweezer</td>
			<td>Stoelting, USA</td>
			<td>52108-83P</td>
			<td></td>
		</tr>
		<tr>
			<td>PINNACLE system</td>
			<td>Pinnacle Technology, USA</td>
			<td>8400-K3-SL</td>
			<td>System for recording EEG (2 channels) and EMG (1 channel) of mice</td>
		</tr>
		<tr>
			<td>Shaving machine</td>
			<td>Braun</td>
			<td>Series 3310s</td>
			<td></td>
		</tr>
		<tr>
			<td>Single and multi-channel pipettes</td>
			<td>Eppendorf, Austria</td>
			<td>Epp 3120 000.020, Epp 3122 000.019</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Kraspharma, Russia</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Soldering station</td>
			<td>AOYUE, China</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotaxic frame</td>
			<td>Stoelting, USA</td>
			<td>51500</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight dissecting scissors</td>
			<td>Stoelting, USA</td>
			<td>52132-10P</td>
			<td></td>
		</tr>
		<tr>
			<td>Tetracycline</td>
			<td>JSC Tatkhimfarmpreparaty, Russia</td>
			<td>N/A</td>
			<td>Eye ointment</td>
		</tr>
		<tr>
			<td>Tweezer</td>
			<td>Stoelting, USA</td>
			<td>52100-03</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasonic cell disrupter</td>
			<td>Biobase, China</td>
			<td>USD-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Wound retractor</td>
			<td>Stoelting, USA</td>
			<td>52125</td>
			<td></td>
		</tr>
		<tr>
			<td>Xylanit</td>
			<td>Nita-Farm, Russia</td>
			<td>N/A</td>
			<td>Muscle relaxant</td>
		</tr>
		<tr>
			<td>Zoletil 100</td>
			<td>Virbac Sante Animale, France</td>
			<td>N/A</td>
			<td>General anesthesia</td>
		</tr>
	</tbody>
</table>

photobiomodulation under electroencephalographic controls of sleep for stimulation of lymphatic removal of toxins from mouse brain

The meningeal lymphatic vessels (MLVs) play an important role in the removal of toxins from the brain. The development of innovative technologies for the stimulation of MLV functions is a promising direction in the progress of the treatment of various brain diseases associated with MLV abnormalities, including Alzheimer&#39;s and Parkinson&#39;s diseases, brain tumors, traumatic brain injuries, and intracranial hemorrhages. Sleep is a natural state when the brain&#39;s drainage processes are most active. Therefore, stimulation of the brain&#39;s drainage and MLVs during sleep may have the most pronounced therapeutic effects. However, such commercial technologies do not currently exist.
This study presents a new portable technology of transcranial photobiomodulation (tPBM) under electroencephalographic (EEG) control of sleep designed to photo-stimulate removal of toxins (e.g., soluble amyloid beta (A&#946;)) from the brain of aged BALB/c mice with the ability to compare the therapeutic effectiveness of different optical resources. The technology can be used in the natural condition of a home cage without anesthesia, maintaining the motor activity of mice. These data open up new prospects for developing non-invasive and clinically promising photo-technologies for the correction of age-related changes in the MLV functions and brain&#39;s drainage processes and for effectively cleansing brain tissues from metabolites and toxins. This technology is intended both for preclinical studies of the functions of the sleeping brain and for developing clinically relevant treatments for sleep-related brain diseases.

Author Spotlight: Developing Photo-Stimulation Technology for Toxin Removal in Sleep-Related Brain Diseases

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

In the study we propose a new way of technology for photo-stimulation of many germ fighting vessels and brain drainage doing sleep and control. Sleep is a natural state of activation of drainage and cleansing function of the brain which is reduced in large numbers of diseases of CNS. It's necessary to create technologies for treating brain diseases during sleep in order to most effectively stimulate the cleansing of its tissues from toxins and metabolists with the flow of food.The technology we have developed for photo-biomulation of the Amylin functions in brain drainage opens up new possibilities from non-pharmacological alternative preferred for a wide prevention of diseases of CNS. They are shaded with a special version of brain drainage including brain or cord injuries, Parkinson's disease and multiple sclerosis, diabetes, et cetera.

In the first step, the study has focused on establishing the effective light dose (a 1050 nm LED) for stimulation of lymphatic removal of fluorescent A&#946; from the brain to dcLNs in awake adult (2-3 month old, 26-29 g) male BALB/c mice. The light doses were selected randomly as 10 J/cm2, 20 J/cm2, and 30 J/cm2 based on our previous studies of tPBM effects on the removal of different dyes and the red blood cells from the brain7,<sup class="x...

This study presents an original and portable tPBM technology under electroencephalographic (EEG) control of deep or non-rapid eye movement (NREM) sleep in non-anesthetized male C57BL/6 mice of different ages to stimulate lymphatic clearance of A&#946; from the brain into the peripheral lymphatic system (the deep cervical lymph nodes, dcLNs).

The meningeal lymphatic vessels (MLVs) play an important role in the removal of toxins from the brain. The development of innovative technologies for the ...

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Research

JoVE Journal

Neuroscience

329 Views.  Saratov State University. In the study we propose a new way of technology for photo-stimulation of many germ fighting vessels and brain drainage doing sleep and control. Sleep is a natural state of activation of drainage and cleansing function of the brain which is reduced in large numbers of diseases of CNS. It's necessary to create technologies for treating brain diseases during sleep in order to most effectively stimulate the cleansing of its tissues from toxins and metabolists with the flow of food.The tech...

Video: Author Spotlight: Developing Photo-Stimulation Technology for Toxin Removal in Sleep-Related Brain Diseases

Hardware Assembly to Perform Transcranial Photobiomodulation for Brain Drainage in Aged Mice

To begin the hardware assembly, solder the light emitting diode or LED to the printed circuit board or PCB, and place the LED and magnets on the 3D frame. Then connect the resistor between an LED anode and the five volt port on the Arduino. Next, connect an LED cathode to the metal oxide semiconductor field effect transistor or MOSFET drain, and secure the MOSFET source to the ground.After connecting the pull down resistor between the MOSFET gate and the ground, proceed to pin the MOSFET gate to pin three on the Arduino. Then connect a liquid crystal display or LCD keypad shield to the Arduino. And insert the Arduino board with LCD keypad shield MOSFET and LED connector into the case.Finally, download the Arduino sketch and open it via the Arduino integrated development environment. Select the correct communication port and flash the firmware.

EEG and Catheter Implantation in Mouse Brain for Transcranial Photobiomodulation to Stimulate Meningeal Lymphatic Vessel Functions

Place the anesthetized mouse in a stereotaxic frame over a heating pad and apply ophthalmic ointment to the eyelids to prevent the eyeballs from drying. Shave the head in the area from the nasal bones to the occipital bones, and disinfect the exposed skin with alternating rounds of chlorhexidine and alcohol three times each. Using straight dissecting scissors, cut off the scalp, hold it with micro forceps, and clean the skull from fascia.Using a drill with a 1.3 millimeter diameter, make two holes in the temporal bones on each side along the coordinates for the two screw pairs. Place the electroencephalographic, or EEG screws, with wire leads in alcohol for 15 minutes, followed by the saline solution. Place four silver plated screws with electrodes into the holes to a depth of one millimeter.Fix the screws on the surface of the skull using dental acrylic, followed by attaching an EEG recording sensor to the animal's nose. After 30 minutes, use curved tweezers to fix the EMG electrodes on the back of the orbicularis oculi muscle with dental acrylic. Connect and fix the EEG electrodes to the silver-plated recess of the sensor.Finally, place the mouse on a heating pad to maintain body temperature until the animal fully recovers from anesthesia. After seven days, use dental acrylic and Dumont forceps to fix a metal plate with a diameter of five millimeters on the occipital bone of the mouse skull. To prepare the chronic catheter, mark a segment on the insulin needle two millimeters from the side of its beveled end.Fix the insulin needle in the holder from the side of the beveled tip to the marked segment. Place a two-centimeter PE-10 polyethylene catheter over the entire remaining length of the needle. Fill the catheter with a saline solution and cover it with a plastic cap.For the implantation, drill a 1.5 millimeter diameter trepanation hole at the appropriate coordinates. Place the PE-10 polyethylene catheter in a stereotactic holder and insert it into the mouse's skull. Afterward, fix it with dental acrylic and let it harden for 15 minutes.Connect any commercial EEG recording system to the connector on the mouse's head, and set the requirement value of the pulse width modulation, or PWM duty cycle. Monitor the EEG signal and wait for delta rhythm activity. If non-rapid eye movement, or NREM, sleep is seen, initiate the photobiomodulation, or PBM process and stop the process if NREM sleep transitions to rapid eye movement sleep or wakefulness.

Confocal Imaging to Assess Lymphatic Removal of Beta-Amyloid from Mouse Brain After Transcranial Photobiomodulation

For confocal imaging, first connect a 10-centimeter catheter to an insulin needle. Using a Hamilton syringe with a 29-gauge needle, prepare five microliters of fluorescent beta amyloid for infusion into the catheter. After fixing the mouse's hand, connect the catheter through a needle to the implanted chronic catheter.Connect the catheter to a micro injector and place the mouse in an individual box. Set the injection rate to 0.1 microliters per minute in the micro injector menu, and press the Start button to inject FA beta into the right lateral ventricle. After FA beta administration, make photobiomodulation, or PBM, using an LED for 61 minutes following the algorithm.Then intravenously inject any tracer for labeling the cerebral vessels via the tail. Post euthanization, use sharp straight scissors to make a small transverse incision in the skin along the trachea, holding the skin with straight non-sharp tweezers. Then with straight scissors, make a longitudinal incision along the entire length of the neck.Employing 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 to push back the surrounding tissues. Make use of two curved tweezers from both sides of the neck to examine the area between the trachea and the cleidomastoid muscle.With straight tweezers having blunt ends, take off the deep cervical lymph node on each side and cut it from the connective tissue. Finally, place the nodes in a Petri dish with a saline solution and cover them with horizontally oriented cover glass to perform confocal microscopy. Light doses of 30 joules per square centimeter showed the most significant effect for promoting lymphatic removal of FA beta from the brains of awake adult male mice.The pulse mode at 1050 nanometers proved most effective in enhancing the removal of FA beta compared to other wavelengths and the continuous mode at the same wavelength. A 10-day course of transcranial PBM during NREM sleep significantly reduced soluble FA beta levels in aged mice, aligning them with levels found in younger adult mice. This effect was not observed when the treatment was applied during wakefulness.