LH was funded by NPL&#39;s MedAccel program financed by the Department for Business, Energy and Industrial Strategy&#39;s Industrial Strategy Challenge Fund. JMG received funding from the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) under project GR 5824/1. JJ acknowledges the funding support from Academy of Medical Sciences Springboard (REF: SBF007\100007) award. SEB acknowledges support from Cancer Research UK under grant number C9545/A29580. AMI, BZ, and SR were supported by U.K. Department for Business, Energy &#38; Industrial Strategy via funding of the National Measurement System. Figure 1 and Figure 2 were created with BioRender.

Table 1: Overview of acoustic and optical properties found in soft tissues. Optical properties cover a spectrum ranging from 600 to 900 nm. It should be noted that these are only representative values intended to provide general guidance. Precise values may vary depending on the experimental condition (e.g., temperature) and frequency/wavelength. The literature provides more specific values. *No specific reference found. <a href="https://www.jove.com/files/ftp_upload/65475/65475_Table 1_rev.xlsx" target="_blank">Please click here to download this Table.</a>
The given protocol has been developed for making ~120 mL of phantom material. The masses of the components can be scaled to make different volumes of phantom material. Please note that for larger volumes (&#62;500 mL), the proposed equipment may not be able to sufficiently heat the phantom mixture homogenously. For this purpose, heating equipment should be adapted appropriately.
CAUTION: Always ensure suitable personal protective equipment (PPE) is worn throughout the entire fabrication process. This may include the use of lab coats, safety goggles, and safety gloves; refer and adhere to local safety guidelines. The procedure is adapted from Hacker et al.39; a summary of the steps is displayed in Figure 1.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65475/65475fig01.jpg" /> 
Figure 1: Fabrication of the copolymer-in-oil material. (1) Materials for optical scattering and absorption are added to mineral oil and (2) sonicated at 90 &#176;C until dissolved. (3) Polymer(s) and stabilizer are added, and (4) the mixture is heated up in an oil bath to 160 &#176;C under low stirring. (5) Upon dissolution of all the components, the sample is poured into a suitable phantom mold and (6) left to harden at room temperature. This figure is reproduced from Hacker et al.39. <a href="https://www.jove.com/files/ftp_upload/65475/65475fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
1. Preparation of the mixture
<ol>
	<li>Prepare the Nigrosin stock solution by adding 0.4 g of Nigrosin to 40 mL of mineral oil. Ensure homogeneous mixing by sonication and thorough vortexing of the sample. Store the stock solution at room temperature. 
	NOTE: The stock solution should always be thoroughly mixed before reuse. If a phantom material without an optical scatterer (TiO2) or absorber (dye) is preferred, step 1 and 2&#160;can be skipped. Proceed with step 3.</li>
	<li>Sonicate 0.15 g of TiO2 and 1 mL of the dye stock solution in 100 mL (83.8 g) of mineral oil until all the components have been completely dissolved (~60 min) (Figure 1: steps 1 and 2). Set the sonicator to elevated temperatures (90 &#176;C) if the equipment allows, as this facilitates the mixing process. Proceed with steps 3-5 during the sonication time. 
	NOTE: If a phantom with higher absorbing and scattering properties is preferred, the sonication time may need to be extended.</li>
	<li>Weigh out the polystyrene-block-poly(ethylene-ran-butylene)-block-polystyrene (SEBS) and low-density polyethylene (LDPE) at the desired concentrations (e.g., SEBS = 25.14 g; LDPE = 6.70 g) (Figure 1: step 3).
	<ol>
		<li>Optional: An antioxidant may be added to increase stability, but it is not obligatory if the heating temperatures do not exceed 180&#176;C. 
		NOTE: In case solubility or viscosity issues are encountered at a later stage of the manufacturing process, it is advisable to exclude the LDPE. LDPE is incorporated to enhance the speed of sound in the material (Table 3); however, it is not mandatory for creating a stable phantom. By omitting LDPE, the manufacturing and molding process can be simplified, but it will result in a decrease in the subsequent speed of sound of the final material (Table 3).</li>
	</ol>
	</li>
	<li>Create an oil bath using suitable glassware and silicone oil; carefully secure it on the hotplate. Ensure the thermocouple remains in the silicone oil bath and does not touch the edges of the glassware throughout the procedure (Figure 2). 
	NOTE: Ensure that the thermoregulatory accessory is carefully mounted, as instructed by the equipment manufacturer.</li>
	<li>Place a magnetic stir bar of adequate length inside the oil bath to ensure uniform heat distribution.</li>
	<li>Turn on the hotplate, set the heating temperature to 160 &#176;C, and set the revolutions per minute (rpm) of the stirrer to 50.</li>
	<li>Transfer the LDPE and SEBS into the glass beaker containing the sonicated mineral oil (with TiO2 and Nigrosin). Introduce a magnetic stir bar of adequate length into the glass beaker and transfer it into the center of the oil bath for heating of the measured components. Ensure that the oil level in the bath remains above the mineral oil level inside the beaker (Figure 1: step 4).</li>
</ol>
2. Heating the mixture
<ol>
	<li>If at any stage the added polymer appears to float over the mineral oil, manually stir the mineral oil solution using a metallic spatula, so that any floating polymer is distributed inside the mineral oil. Wear heat-resistant gloves.</li>
	<li>Leave the mixture at 160 &#176;C until all the polymer has dissolved and the solution appears uniformly mixed, with a smooth and homogeneous texture (~1.5 h).</li>
</ol>
3. Vacuuming
NOTE: For removal of air bubbles, follow the following steps, depending on the equipment available.
<ol>
	<li>Place the hot beaker carefully into the vacuum chamber and vacuum the samples for 2-3 min on the highest setting (lowest vacuum). Use a metallic spatula to carefully remove any air bubbles that accumulate on the surface. If air bubbles are still present after this step, reheat the mixture and repeat the vacuuming step until all air bubbles have been removed.</li>
	<li>Turn on the vacuum oven and heat it up to 160 &#176;C. Once it has reached the desired temperature, transfer the beaker with the solution into the vacuum oven. 
	NOTE: Always handle the beaker with heat protective gloves.
	<ol>
		<li>Switch on the vacuum to the highest setting (lowest vacuum) available. If a foam layer has produced on top of the solution, turn off the vacuum and remove the bubbles from the surface using a spatula (repeat this step until all air bubbles have been removed).</li>
		<li>Leave the beaker in the vacuum oven for 1 h at the highest vacuum setting. 
		&#8203;NOTE: To keep the vacuum oven clean, clean the external surface of the beaker of silicone oil with a paper towel. </li>
	</ol>
	</li>
</ol>
4. Pouring the samples into the mold
<ol>
	<li>Before pouring the samples into sample molds, remove any remaining air bubbles on the surface of the mixture with a spatula if necessary.</li>
	<li>Carefully pour the solution into a suitable mold, wearing heat-resistant gloves or using adequate protective equipment. Ensure smooth and steady pouring from a low height to reduce the chance of any air bubbles forming (Figure 1: step 5). For molds with complex shapes, coat the mold with a thin layer of oil (other than mineral oil [e.g., castor or silicone oil]) prior to pouring to facilitate removal of the cured sample. 
	NOTE: Preheating of the molds in an oven can help with achieving higher sample homogeneity.</li>
	<li>Remove any air bubbles from the top of the samples quickly with a metallic spatula once poured. If numerous air bubbles have accumulated within the mixture, repeat the vacuum step, provided that the type and shape of the mold permits it.</li>
	<li>Allow the solution to set at room temperature. Although smaller samples may cure in less than 2 h, leave the samples overnight to eliminate any risk of incomplete curing. Store the samples at room temperature (Figure 1: step 6).</li>
</ol>
5. Image acquisition
<ol>
	<li>For image acquisition, place the phantom in the field of view of the imaging device.</li>
	<li>For PAI or ultrasound systems, perform acoustic coupling of the phantom surface to the ultrasound transducer, for example, with ultrasound gel or water. 
	NOTE: If the protocol has been followed correctly, no inhomogeneities should perturb the field of view. Custom phantom holders can assist in repeatable sample positioning between measurements.</li>
	<li>If the acquisition temperature differs from the storing temperature of the phantom, allow the phantom temperature to stabilize with the surroundings.</li>
	<li>Acquire an image.</li>
</ol>
6. Material characterization measurements
NOTE: The purpose of material characterization measurements is verification of the material optical and acoustic properties. It should be noted that the phantom fabrication protocol has shown high reproducibility39, so the general measurement protocols that follow are only provided as guidance if further verification studies are desired. The individual steps of the measurements will depend on the characterization equipment used. Here, a system based on a broadband through-transmission substitution method41 (available at the National Physical Laboratory (NPL), UK) was employed for acoustic characterization and an in-house double-integrating-sphere (DIS) system was used (based on 42) for optical characterization. The setup of the characterization systems is displayed in Supplementary Figure 1. Additional details on the measurement setups (acoustic43; optical42,44) and measurement procedure39 can be found elsewhere. The measurement procedure should be adapted accordingly to each specific characterization system used.
<ol>
	<li>Acoustic characterization 
	NOTE: The acoustic characterization measurements are based on a system employing a 10 MHz center frequency ultrasound transducer (active element diameter of 10 mm) for pulse generation and a broadband hydrophone (30 mm active element diameter bilaminar membrane hydrophone) for pulse detection (both placed in a water tank filled with deionized water; dimensions of 112 cm x 38 cm x 30 cm3). The transducer is driven by a pulser-receiver. Waveforms are acquired using an oscilloscope. More details on the setup and measurement procedure (including system-specific type B effects on the measurements) can be found in 43.
	<ol>
		<li>Prepare samples suitable for the measurement setup (e.g., in this case, circular samples with a diameter of 7-8 cm and thickness of 6-9 mm). Ensure that the samples are of homogeneous composition and free of any impurities, air bubbles, or surface irregularities.</li>
		<li>Measure the thickness of the test sample using vernier calipers and record the temperature of the water tank using a calibrated thermometer.</li>
		<li>Place the sample into the system. Ensure that the sample is correctly aligned with the components of the system. 
		NOTE: An automatically controlled sample holder based on a gimbal mount43 may assist in precise control of the rotation and tilt of the sample.</li>
		<li>Acquire four acoustic pulses for each measurement set: a reference through-water pulse with no sample present in the acoustic path; a through-sample transmission; and acoustic reflections received at the transmitter from the front and rear surfaces of the sample.</li>
		<li>Derive the acoustic properties of the sample from the measurements. Calculate the speed of sound c(f) (in m&#183;s&#8722;1) using the equation (1)43. 
		<img alt="Equation 1" src="/files/ftp_upload/65475/65475eq01.jpg" style="margin: auto;" />&#160; &#160; (1) 
		cw depicts the temperature-dependent speed of sound of water, and &#952;1(f), &#952;2(f), &#952;w(f), and &#952;s(f) are the corresponding unwrapped phase spectra of the front-reflected, back-reflected, through-water, and through-sample voltage pulses, respectively. The frequency-dependent attenuation coefficient (&#945;i(f)) of the material can be derived using the two-sample substitution technique shown in equation (2)43. 
		<img alt="Equation 2" src="/files/ftp_upload/65475/65475eq02v2.jpg" style="margin: auto;" />&#160;&#160;&#160; (2) 
		Uw(f) and Us(f) are the respective voltage magnitude spectra of the through-water and through-sample pulse, &#945;w(f) is the attenuation coefficient of ultrasound (in dB&#183;cm-1) of pure water at the specific water tank temperature45, and d1 and d2 (d2 &#62; d1) are two sample thicknesses. 
		NOTE: For the present protocol, the attenuation accounting for interfacial losses was assessed, finding its impact to be negligible.</li>
		<li>Repeat the measurement more than three times at different positions on the test sample. Calculate the mean and standard deviation of the measurements to derive a final sample value.</li>
	</ol>
	</li>
	<li>Optical characterization 
	NOTE: For optical testing, a double-integrating sphere system was used (based on 42), employing two integrating spheres (50 mm internal diameter) that are connected to two spectrometers via two optical fibers. The reflectance sphere is connected to a light source via a third optical fiber.
	<ol>
		<li>Prepare samples suitable for the measurement setup (e.g., in this case, rectangular samples with a width of 5.9 cm, height of 1.8 cm, and thickness ranging between 2 and 3 mm). Ensure that the samples are of homogeneous composition and free of any impurities, air bubbles, or surface irregularities.</li>
		<li>Turn on the light source and allow it to stabilize according to the manufacturer&#39;s instructions (e.g., 15 min).</li>
		<li>Determine the thickness of the sample using vernier calipers. If applicable, specify the wavelength range and step size for the measurement (e.g., 450-900 nm with a 1 nm step size).</li>
		<li>Record the reference measurements for the transmission and reflectance sphere.
		<ol>
			<li>For the reflectance sphere, first take an open port measurement by recording the reflectance value&#160;R0 with the transmittance sphere removed and light source turned on. Then, record the reflectance value R1 with a reference standard held in front of the reflectance sphere (light source turned on).</li>
			<li>For the transmittance sphere, first take a blocked beam measurement by recording the transmittance value T0 with the aligned reflectance and transmittance spheres and light source turned off. Then, take an incident beam measurement by recording the transmittance value T1 with the aligned reflectance and transmittance spheres and light source turned on. 
			NOTE: A clean surface of the spheres and reference standard must be ensured for the measurements, as the adherence of dust or other contaminants may impact the performance of the components46.</li>
		</ol>
		</li>
		<li>Place the sample between the spheres. Measure the reflectance Rs and transmittance Ts&#160;values. Ensure that the sample is not compressed, as this may impact the measurement accuracy. Placing one sphere on a motorized stage may help to accurately control the distance between the spheres by adapting it to the measured sample thickness.</li>
		<li>Calculate the normalized reflectance MR&#160;and transmittance MT&#160;values using equations (3) and (4)42. 
		<img alt="Equation 3" src="/files/ftp_upload/65475/65475eq03.jpg" style="margin: auto;" />&#160; &#160; (3) 
		<img alt="Equation 4" src="/files/ftp_upload/65475/65475eq04.jpg" style="margin: auto;" />&#160; &#160; (4) 
		rstd depicts the intensity reflected from the 99% reflectance standard.</li>
		<li>Enter the measured values into an inverse adding doubling (IAD) program (source code: http://omlc.org/software/iad/)44 to estimate the optical properties of the material. 
		NOTE: Based on previous reports, the scattering anisotropy factor (g) can be taken as g = 0.7, and the refractive index as n = 1.4 30.</li>
		<li>Repeat the measurement at least three times at different positions along the test sample. Calculate the mean and standard deviation of the measurements to derive a final sample value.</li>
	</ol>
	</li>
</ol>

Fabrication of a Stable, Biologically Relevant Phantom Material

Sarah Bohndiek has previously received research support from CYBERDYNE INC and iThera Medical GmbH, vendors of photoacoustic imaging instruments. The other authors have no conflict of interest related to the present manuscript to disclose.

Here, a protocol is presented that aims to provide a versatile recipe for a stable, biologically relevant material that can be used to create phantoms for calibration measurements and standardization across multimodal acoustic and optical biomedical imaging applications. The material has previously been shown to be stable over time39, to have high batch-to-batch reproducibility, to be safe to use, and to consist of readily available, cost-effective ingredients from standard scientific suppliers. The material properties are independently tunable across relevant acoustic and optical regimes. Furthermore, it is mechanically robust and insoluble in water, thereby withstanding rough handling, and it is inert to water-based coupling agents that are used in ultrasonic/photoacoustic research. It was highlighted that different phantom designs can be created with different types of inclusions, composed of the same or of different material types. Given these properties, the material fulfills the aforementioned key criteria for an ideal biophotonic phantom and shows key advantages compared to other existing tissue-mimicking materials3. By detailing the exact manufacturing process, we hope to minimize variations arising from the fabrication procedure, thereby optimizing its use for calibrating, validating, and tracking the performance of imaging systems.
Two key steps have been identified as being critical to the fabrication process. First, ingredients need to be thoroughly mixed and uniformly heated for the creation of a homogenous material. Using a sonicator and magnetic stirrer for mixing and an oil bath for heating ensures the even distribution of material components within the base matrix. Care needs to be taken that the oil bath does not reach very high temperatures (&#62;180 &#176;C), as this will result in oxidation of the material components, leading to yellowish discoloration. Manual stirring can support the mixing process and compensates for insufficient heating from the material-air interface. The time for sonication and mixing may need to be extended when a higher concentration of TiO2 and/or polymers are used to ensure a homogenous composition of the material. Second, air bubbles need to be removed to prevent the formation of heterogeneities within the base matrix. Whilst this can be achieved with a vacuum pump or oven, careful pouring from a low height should also be practiced to minimize trapping air within the material.
One significant advantage of the material is its thermoplastic properties (derived from the SEBS polymer), allowing it to be reheated and remolded without any significant impact on its acoustic and optical properties39. However, reheating needs to be performed gradually and carefully, as the material can easily burn and oxidize if reheated too quickly. Reheating also becomes more difficult when higher LDPE concentrations are used, as LDPE does not exhibit the same thermoplastic behavior as SEBS.
Several limitations of the protocol remain. Due to the high melting temperature of the polymers (150 &#176;C), phantom molds need to made out of a heat-resistant material, such as glass or stainless steel. Additionally, the material is fairly viscous in liquid state if a high polymer concentration is used to tune the acoustic properties, making the filling of small imaging targets difficult. Finally, tuning of the acoustic properties is thus far limited to a speed of sound range of ~1450-1,516 m&#183;s-1 which supports mimicking tissues such as breast or fat (c = 1,450-1,480 m&#183;s&#8722;1), but may be insufficient for tissues like muscle or kidney (c &#62; 1,520 m&#183;s&#8722;1). The concomitant change of acoustic attenuation should also be taken into consideration.
Here, we have highlighted the application of the material as a stable phantom for ultrasound and optical imaging applications. However, copolymer-in-oil materials have also shown to be of value in elastography applications35, and could potentially allow for compatibility with further imaging modalities such as magnetic resonance imaging. Increased anatomical realism of the phantoms may be achieved using 3D-printed molds, as shown in similar studies29,47,48,49. Early studies have also demonstrated the 3D printability of the material itself, further extending its flexibility in terms of processing and fabrication. These developments highlight the exciting future potential of the material as a widely used, stable phantom medium for multimodal imaging applications.

Establishing the precision and accuracy of novel optical imaging biomarkers through technical validation1,2 is paramount to ensuring their successful implementation in clinical practice. To achieve this, technical validation studies frequently employ durable physical phantoms, which facilitate inter-instrument performance assessment and routine quality control. For widespread use of a phantom material in research and clinical translation, a simple, highly reproducible fabrication protocol is required. An ideal biophotonic phantom material should include the following properties3: (1) independently tunable properties within biologically relevant ranges; (2) mechanical robustness; (3) long-term stability; (4) flexibility in geometry and architecture; (5) safe handling; (6) widely available ingredients that can be purchased from standard scientific suppliers; and (7) low cost. At present, biophotonic applications lack a standardized protocol for a widely accepted phantom material that fulfils the outlined requirements and also includes tunable acoustic properties for hybrid applications, such as photoacoustic imaging (PAI).
Biologically relevant phantom materials targeted for combined optical and acoustic applications include hydrogels4,5, polyvinyl alcohol (PVA)6,7,8,9, and polyvinyl chloride plastisol (PVCP)10,11,12,13,14,15,16. However, these materials are characterized by certain limitations that restrict their application as a stable phantom material. Hydrogels, for example, are prone to dehydration, mechanical damage, and bacterial ingrowth, limiting their shelf life17,18,19. The addition of chemicals can increase the longevity, but common preservatives, such as formaldehyde20 or benzalkonium chloride21, are hazardous and require cautionary measures during handling. Additionally, targets containing water-soluble dyes can diffuse within the base material if not encapsulated. PVA cryogels are characterized by a higher longevity and structural robustness, but their preparation process involves long freeze-thaw cycles22. This can limit the independent tunability of acoustic and optical parameters23 and-if slightly varied-can lead to inhomogeneities6, thereby compromising reproducibility. Moreover, the diffusion of dyes from inclusions has been observed after 1 year13. PVCP has a complex fabrication process that includes high temperatures of up to 180-220 &#176;C13,14,24,25. PVCP also suffers from a lack of a supply chain with scientific suppliers26 and can contain plasticizers based on phthalates, which may cause reproductive and developmental harm27, making them controlled substances in some countries.
Copolymer-in-oil compositions, such as gel wax28,29,30,31 or blends based on thermoplastic styrenic elastomers32,33,34,35,36, exhibit good longitudinal stability and feature tissue-like acoustic and optical properties31,35,36,37, thereby having high potential as a durable phantom candidate in multimodal applications. Additionally, this class of material is cost-effective, non-water absorbing, non-toxic, and biologically inert35,38. The speed of sound c(f) and acoustic attenuation coefficient &#945;(f) can be tuned in a biologically relevant range (Table 1) by variation of the polymer concentration33,35,39, whilst optical absorption &#181;a(&#955;) and reduced scattering &#181;s&#39;(&#955;) coefficients can be primarily varied by the addition of oil-soluble dyes or titanium dioxide (TiO2)39, respectively.
Here, a simple, easy-to-follow protocol is presented for the creation of durable copolymer-in-oil phantoms suitable for use in optical, ultrasound, or photoacoustic device calibration. All ingredients have defined Chemical Abstract Service (CAS) numbers and are readily available from standard scientific suppliers. Potential difficulties in the fabrication procedure are highlighted and ways to overcome them are presented. Whilst the protocol allows the fabrication of materials with a range of acoustic and optical properties, the presented protocol yields a material with a speed of sound of ~1,481 m&#183;s-1, aligning with the speed of sound of water at room temperature (20 &#176;C)40. This value was selected as a neutral standard for representing the wide range of existing tissue properties (Table 1), allowing the establishment of a consistent and reliable reference point for comparison. By providing this detailed protocol, we aim to broaden the uptake and manufacturing reproducibility of this promising phantom material type, thereby facilitating biophotonic, acoustic, and photoacoustic validation studies and supporting routine quality control in preclinical and clinical imaging applications.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Low-density Polyethylene (LDPE)</td>
			<td>Alfa Aesar&#160;</td>
			<td>43949.30 
			CAS: 9002-88-4</td>
			<td>6.70 g 
			5.8% w/w</td>
		</tr>
		<tr>
			<td>Mineral oil</td>
			<td>Sigma Aldrich</td>
			<td>330779-1L 
			CAS: 8042-47-5</td>
			<td>83.80 g 
			72.5% w/w</td>
		</tr>
		<tr>
			<td>Nigrosin&#160;</td>
			<td>Sigma Aldrich&#160;</td>
			<td>211680-100G 
			CAS: 11099-03-9</td>
			<td>0.4 g&#160;</td>
		</tr>
		<tr>
			<td>Polystyrene-block-poly(ethylene-ran-butylene)-block-polystyrene (SEBS)</td>
			<td>Sigma Aldrich&#160;</td>
			<td>200557-250G 
			CAS: 66070-58-4</td>
			<td>25.14 g 
			21.7% w/w</td>
		</tr>
		<tr>
			<td>Silicone oil for oil bath</td>
			<td>Sigma Aldrich&#160;</td>
			<td>85409-1L 
			CAS: 63148-62-9 
			~1 L (depending on size of oil bath)</td>
		</tr>
		<tr>
			<td>Titanium(IV) oxide, anatase (TiO2)</td>
			<td>Sigma Aldrich&#160;</td>
			<td>232033 
			CAS: 1317-70-0</td>
			<td>0.15 g</td>
		</tr>
		<tr>
			<td>Antioxidant (optional; e.g., butylated hydroxytoluene)</td>
			<td>Sigma Aldrich&#160;</td>
			<td>W218405-1KG-K 
			CAS: 128-37-0</td>
			<td>may be added to prevent discolouration if higher fabrication temperatures are used&#160;</td>
		</tr>
		<tr>
			<td>Oil-solule dyes for inclusions (optional)</td>
			<td>e.g., Caligo safe wash relief inks, Cranfield Colours, Cwmbran, UK*</td>
			<td></td>
			<td>may be added depending on preferred absorption</td>
		</tr>
		<tr>
			<td>Bath Sonicator&#160;</td>
			<td>Ultrawave Ltd, UK*</td>
			<td>U500H Ultrasonic Cleaning Bath*</td>
			<td>ideally with temperature control; recommended operating frequencies between 30-50 kHz)</td>
		</tr>
		<tr>
			<td>Crystallising dish for silicone oil bath (up to +200 &#176;C)</td>
			<td>any suitable supplier</td>
			<td></td>
			<td>(ext. diameter ~140 mm for 250 mL glass beaker)</td>
		</tr>
		<tr>
			<td>Glass beaker (~250 mL); glass sample dishes/phantom mould</td>
			<td>any suitable supplier</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hot plate with magnetic stirrer and thermoregulator (external probe thermocouple) with maximum temperature of &#62; 200 &#176;C</td>
			<td>Thermo Fisher Scientific, UK*</td>
			<td>Velp Scientifica AREC.X Digital Ceramic Hot Plate Stirrer*</td>
			<td></td>
		</tr>
		<tr>
			<td>Laboratory scales / scientific balances accurate to 0.01 g</td>
			<td>any suitable supplier</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Metallic spatula</td>
			<td>any suitable supplier</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Vaccuum oven or vaccum chamber</td>
			<td>Memmert, Germany (Vacuum oven)* 
			 
			Cole-Parmer Instrument Company, UK (Vaccum chamber)*</td>
			<td>VO29 (Vacuum oven) 
			 
			DWK Life Sciences (Kimble) Glass Vacuum Desiccator with Collar (Vaccum chamber)*</td>
			<td></td>
		</tr>
		<tr>
			<td>1. Acoustic characterization system*</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrophone</td>
			<td>GEC Marconi</td>
			<td>30 mm active element diameter bilaminar membrane hydrophone</td>
			<td></td>
		</tr>
		<tr>
			<td>Oscilloscope</td>
			<td>Tektronix UK, Bracknell, UK</td>
			<td>DPO 7254&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Pulser&#8211;receiver&#160;</td>
			<td>Olympus NDT, Waltham, MA, USA</td>
			<td>Olympus 5073PR</td>
			<td></td>
		</tr>
		<tr>
			<td>Sample holder</td>
			<td>Newport Spectra-Physics, Didcot, UK</td>
			<td>Newport 605-4 Series Low-Distortion Axial Clamping Gimbal Optic Mount</td>
			<td>may require additional adaptor for sample holding</td>
		</tr>
		<tr>
			<td>Thermometer</td>
			<td>G. H. Zeal, London, UK</td>
			<td>UKAS-calibrated IP 39C spirit-in-glass thermometer</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasound transducer&#160;</td>
			<td>Force Technology, Brondby, Denmark</td>
			<td>Transducer of active element diameter 10 mm&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Vernier callipers</td>
			<td>any suitable supplier</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Water tank filled with deionized water&#160;</td>
			<td>any suitable supplier</td>
			<td></td>
			<td>requires sufficent size (e.g., dimensions 112 &#215; 38 &#215; 30 cm3)</td>
		</tr>
		<tr>
			<td>2.Optical characterization system*</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Integrating sphere (two)</td>
			<td>Avantes, Apeldoorn, the Netherlands</td>
			<td>AvaSphere-50, 50 cm internal diameter</td>
			<td></td>
		</tr>
		<tr>
			<td>Light source&#160;</td>
			<td>Avantes, Apeldoorn, the Netherlands</td>
			<td>Avalight-HAL-s-mini</td>
			<td></td>
		</tr>
		<tr>
			<td>Motorized stage (optional)</td>
			<td>Thorlabs,</td>
			<td>Thorlabs MTS50</td>
			<td></td>
		</tr>
		<tr>
			<td>Optical fibres (three)</td>
			<td>any suitable supplier</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Reflectance standard</td>
			<td>Labsphere, North Sutton, USA</td>
			<td>99%, white, 1.25&#34; diameter, USRS-99-010, AS-01158-060</td>
			<td></td>
		</tr>
		<tr>
			<td>Spectrometer</td>
			<td>Avantes, Apeldoorn, the Netherlands</td>
			<td>Starline Avaspec-2048</td>
			<td></td>
		</tr>
		<tr>
			<td>Software</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Data acqusition software (e.g., Labview)</td>
			<td>National Instruments, Austin, TX, USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Data analysis software (e.g., Matlab)</td>
			<td>Mathworks, Natick, USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Inverse adding doubling (IAD) program&#160;</td>
			<td></td>
			<td></td>
			<td>Source code: http://omlc.org/software/iad/</td>
		</tr>
		<tr>
			<td>*Please note that similar equipment may also be used.</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

a stable phantom material for optical and acoustic imaging

Establishing tissue-mimicking biophotonic phantom materials that provide long-term stability are imperative to enable the comparison of biomedical imaging devices across vendors and institutions, support the development of internationally recognized standards, and assist the clinical translation of novel technologies. Here, a manufacturing process is presented that results in a stable, low-cost, tissue-mimicking copolymer-in-oil material for use in photoacoustic, optical, and ultrasound standardization efforts.
The base material consists of mineral oil and a copolymer with defined Chemical Abstract Service (CAS) numbers. The protocol presented here yields a representative material with a speed of sound c(f) = 1,481 &#177; 0.4 m&#183;s-1 at 5 MHz (corresponds to the speed of sound of water at 20 &#176;C), acoustic attenuation &#945;(f) = 6.1 &#177; 0.06 dB&#183;cm-1 at 5 MHz, optical absorption &#181;a(&#955;) = 0.05 &#177; 0.005 mm-1 at 800 nm, and optical scattering &#181;s&#39;(&#955;) = 1 &#177; 0.1 mm-1 at 800 nm. The material allows independent tuning of the acoustic and optical properties by respectively varying the polymer concentration or light scattering (titanium dioxide) and absorbing agents (oil-soluble dye). The fabrication of different phantom designs is displayed and the homogeneity of the resulting test objects is confirmed using photoacoustic imaging.
Due to its facile, repeatable fabrication process and durability, as well as its biologically relevant properties, the material recipe has high promise in multimodal acoustic-optical standardization initiatives.

Following this recipe, three representative phantom designs have been created for the purpose of photoacoustic imaging, targeted for different system designs with different optical illumination and acoustic detection geometries (Figure 3A). If the phantom preparation procedure is performed successfully, the phantom material appears smooth and homogeneous without any trapped air bubbles or impurities, and no artifacts can be seen in the resulting image (here visualized using photoacoustic imaging; Figure 3B,C). The protocol yields a representative material with a speed of sound c(f) = 1481 &#177; 0.4 m&#183;s-1 (corresponding to the speed of sound of water at 20 &#176;C40), acoustic attenuation &#945;(f) = 6.1 &#177; 0.06 dB&#183;cm-1 (both at 5 MHz), optical absorption &#181;a(&#955;) = 0.05 &#177; 0.005 mm-1, and optical scattering &#181;s&#39;(&#955;) = 1 &#177; 0.1 mm-1 (both at 800 nm) (uncertainty depicts the standard deviation from n = 3 independently produced batches by different operators; all measurements were carried out at room temperature [20 &#176;C]).
The optical scattering coefficient can be tuned by the variation of TiO2, whilst the optical absorption coefficient can be tuned by the addition of any oil-soluble dye, here demonstrated with Nigrosin (Table 2 and Figure 3D). Whilst the values in Table 2 are focused on lower absorbing and scattering tissues, such as muscle or breast (Table 1), we have not encountered any difficulties with adding absorbers and scatterers at higher concentrations. However, the addition of optical scatterers/absorbers at higher concentrations may require longer sonication times to achieve homogenous mixing of the solution.
The acoustic attenuation and speed of sound can be tuned by variation of the polymer concentration (Table 3). Here, tuning is thus far limited to a speed of sound range of ~1,450-1,516 m&#183;s-1. Lower respective polymer concentrations may result in low physical stability of the sample, leading to plastic deformation over time34. Higher polymer concentrations result in brittleness and an uneven texture of the material. The range of acoustic properties may help to mimic tissues such as breast or fat (c = 1,450-1,480 m&#183;s&#8722;1), but may be insufficient for tissues such as muscle or kidney (c &#62; 1,520 m&#183;s&#8722;1; Table 1).
Common error sources in phantom preparation include insufficient removal of air bubbles and inhomogeneous mixing of the base components (Figure 4). This can be minimized by vacuuming and careful pouring, and stirring/vortexing, respectively.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65475/65475fig02.jpg" /> 
Figure 2:&#160;Experimental setup for the phantom fabrication. The glass beaker containing the phantom ingredients is placed in the silicone oil bath using a clamp to avoid direct contact between the surfaces of the oil bath and the glass beaker. Temperature feedback on the hot plate ensures careful temperature control. Magnetic stirrers enable mixing of the both the silicone oil and phantom ingredients. <a href="https://www.jove.com/files/ftp_upload/65475/65475fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65475/65475fig03.jpg" /> 
Figure 3: Representative results from the phantom fabrication procedure. (A) Various phantom designs showing versatility for application in different photoacoustic imaging systems. Left: small rectangular phantom with strings embedded at different depths (0.5, 1.5, and 2.5 mm; inter-target distance of 1.25 mm) designed for testing high-resolution imaging systems; middle: cylindrical phantom with two inclusions (inter-inclusion distance of 12 mm) using a green and violet oil-soluble dye, designed for testing tomography systems; right: large rectangular phantom with channels embedded at different depths (6 mm, 10 mm, and 14 mm; inter-inclusion distance of 3.5 mm), designed for testing a handheld system. (B) Example photoacoustic image of the rectangular phantom with embedded strings, acquired at 532 nm with a commercial photoacoustic imaging system. (C) Example photoacoustic image of the cylindrical tomographic phantom, acquired at 800 nm with a commercial photoacoustic imaging system. (D) Phantoms with increasing optical absorption concentrations by increasing concentrations of Nigrosin (concentrations given in weight percentage of the total volume of mineral oil on the image). Figure 3B,C is reproduced from Hacker et al.39. Scale bars = 10 mm. <a href="https://www.jove.com/files/ftp_upload/65475/65475fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65475/65475fig04.jpg" /> 
Figure 4: Common phantom failures. (A,B) Photographs showing air bubbles trapped inside the base matrix. (C) Insufficient mixing of base components leads to inhomogeneities (red arrows) in the resulting photoacoustic image. Scale bar = 5 mm (A). <a href="https://www.jove.com/files/ftp_upload/65475/65475fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 2: Tabular overview of tuning of optical absorption (&#181;a) and scattering (&#181;s&#39;) values. Percentage values are given as weight percentage to the total volume of the base solution (mineral oil, column 1) and to the total weight of the phantom material (column 2). The Nigrosin concentrations depict the total amount of absolute Nigrosin (not stock solution). All samples included 5% butylated hydroxytoluene as an antioxidant (optional). n = 3 measurements per sample. A visual representation of the table can be found in Hacker et al.39. Abbreviation: neg = negligible. <a href="https://www.jove.com/files/ftp_upload/65475/Table 2v3.xlsx" target="_blank">Please click here to download this Table.</a>
Table 3: Tabular overview of tuning of acoustic attenuation (&#945;) and speed of sound (c) values. Described by the power law &#945;0fn with &#945;0 and n parameters obtained from a non-linear least squares &#64257;tting (n = 4 measurements per sample). F depicts the frequency in MHz. Percentage values are given as weight percentage to the total weight of the base solution (mineral oil). All samples included 5% butylated hydroxytoluene as an antioxidant (optional). A visual representation of the table can be found in Hacker et al.39. <a href="https://www.jove.com/files/ftp_upload/65475/65475_Table 3_rev.xlsx" target="_blank">Please click here to download this Table.</a>
Supplementary Figure S1: Setup of the acoustic and optical characterization systems used for verifications. A photograph (A) and schematic (B) of the acoustic characterization system for determination of the acoustic attenuation coefficient and speed of sound are displayed. Individual system components are denoted by annotations HP (HydroPhone), S (Sample), and T (Transducer) in the photo and schematic. A photograph (C) and schematic (D) of the double-integrating sphere system for the evaluation of the optical absorption coefficient and reduced scattering coefficient are shown. Individual system components are denoted by annotations S (Sample), RS (Reflectance Sphere), TS (Transmission Sphere), OF (Optical Fiber), and MS (Motorized Stage) in the photo and schematic. This figure is reproduced from Hacker et al. 39. <a href="https://www.jove.com/files/ftp_upload/65475/Suppl. Fig 1.pdf" target="_blank">Please click here to download this File.</a>

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Author Spotlight: A Stable Phantom Material for Optical and Acoustic Imaging  

This protocol describes the fabrication of a stable, biologically relevant phantom material for optical and acoustic biomedical imaging applications, featuring independently tunable acoustic and optical properties.

A Stable Phantom Material for Optical and Acoustic Imaging

Establishing tissue-mimicking biophotonic phantom materials that provide long-term stability are imperative to enable the comparison of biomedical imaging ...

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2.5K Views.  University of Cambridge. This protocol describes the fabrication of a stable, biologically relevant phantom material for optical and acoustic biomedical imaging applications, featuring independently tunable acoustic and optical properties. 

Video: Author Spotlight: A Stable Phantom Material for Optical and Acoustic Imaging  

High-Resolution Endocardial and Epicardial Optical Mapping in a Sheep Model of Stretch-Induced Atrial Fibrillation

Atrial fibrillation (AF) is a complex cardiac arrhythmia with high morbidity and mortality.1,2 It is the most common sustained cardiac rhythm disturbance seen in clinical practice and its prevalence is expected to increase in the coming years.3 Increased intra-atrial pressure and dilatation have been long recognized to lead to AF,1,4 which highlights the relevance of using animal models and stretch to study AF dynamics. Understanding the mechanisms underlying AF requires visualization of the cardiac electrical waves with high spatial and temporal resolution. While high-temporal resolution can be achieved by conventional electrical mapping traditionally used in human electrophysiological studies, the small number of intra-atrial electrodes that can be used simultaneously limits the spatial resolution and precludes any detailed tracking of the electrical waves during the arrhythmia. The introduction of optical mapping in the early 90's enabled wide-field characterization of fibrillatory activity together with sub-millimeter spatial resolution in animal models5,6 and led to the identification of rapidly spinning electrical wave patterns (rotors) as the sources of the fibrillatory activity that may occur in the ventricles or the atria.7-9 Using combined time- and frequency-domain analyses of optical mapping it is possible to demonstrate discrete sites of high frequency periodic activity during AF, along with frequency gradients between left and right atrium. The region with fastest rotors activates at the highest frequency and drives the overall arrhythmia.10,11 The waves emanating from such rotor interact with either functional or anatomic obstacles in their path, resulting in the phenomenon of fibrillatory conduction.12 Mapping the endocardial surface of the posterior left atrium (PLA) allows the tracking of AF wave dynamics in the region with the highest rotor frequency. Importantly, the PLA is the region where intracavitary catheter-based ablative procedures are most successful terminating AF in patients,13 which underscores the relevance of studying AF dynamics from the interior of the left atrium. Here we describe a sheep model of acute stretch-induced AF, which resembles some of the characteristics of human paroxysmal AF. Epicardial mapping on the left atrium is complemented with endocardial mapping of the PLA using a dual-channel rigid borescope c-mounted to a CCD camera, which represents the most direct approach to visualize the patterns of activation in the most relevant region for AF maintenance.

This report provides a detailed description of the methodology and results of simultaneous endocardial and epicardial optical mapping of electrical excitation in the intact left atrium of a Langendorff-perfused sheep heart during stretch-induced atrial fibrillation.

 Atrial fibrillation (AF) is a complex cardiac arrhythmia with high morbidity and mortality.1,2 It is the most common sustained cardiac rhythm disturbance ...

Non-invasive Optical Measurement of Cerebral Metabolism and Hemodynamics in Infants

Perinatal brain injury remains a significant cause of infant mortality and morbidity, but there is not yet an effective bedside tool that can accurately screen for brain injury, monitor injury evolution, or assess response to therapy. The energy used by neurons is derived largely from tissue oxidative metabolism, and neural hyperactivity and cell death are reflected by corresponding changes in cerebral oxygen metabolism (CMRO2). Thus, measures of CMRO2 are reflective of neuronal viability and provide critical diagnostic information, making CMRO2 an ideal target for bedside measurement of brain health.
Brain-imaging techniques such as positron emission tomography (PET) and single-photon emission computed tomography (SPECT) yield measures of cerebral glucose and oxygen metabolism, but these techniques require the administration of radionucleotides, so they are used in only the most acute cases.
Continuous-wave near-infrared spectroscopy (CWNIRS) provides non-invasive and non-ionizing radiation measures of hemoglobin oxygen saturation (SO2) as a surrogate for cerebral oxygen consumption. However, SO2 is less than ideal as a surrogate for cerebral oxygen metabolism as it is influenced by both oxygen delivery and consumption. Furthermore, measurements of SO2 are not sensitive enough to detect brain injury hours after the insult 1,2, because oxygen consumption and delivery reach equilibrium after acute transients 3. We investigated the possibility of using more sophisticated NIRS optical methods to quantify cerebral oxygen metabolism at the bedside in healthy and brain-injured newborns. More specifically, we combined the frequency-domain NIRS (FDNIRS) measure of SO2 with the diffuse correlation spectroscopy (DCS) measure of blood flow index (CBFi) to yield an index of CMRO2 (CMRO2i) 4,5.
With the combined FDNIRS/DCS system we are able to quantify cerebral metabolism and hemodynamics. This represents an improvement over CWNIRS for detecting brain health, brain development, and response to therapy in neonates. Moreover, this method adheres to all neonatal intensive care unit (NICU) policies on infection control and institutional policies on laser safety. Future work will seek to integrate the two instruments to reduce acquisition time at the bedside and to implement real-time feedback on data quality to reduce the rate of data rejection.

We combined frequency-domain near-infrared spectroscopy measures of cerebral hemoglobin oxygenation with diffuse correlation spectroscopy measures of cerebral blood flow index to estimate an index of oxygen metabolism. We tested the utility of this measure as a bedside screening tool to evaluate the health and development of the newborn brain.

Perinatal brain injury remains a significant cause of infant mortality  and morbidity, but there is not yet an effective bedside tool that can  accurately ...

In Vivo Optical Imaging of Brain Tumors and Arthritis Using Fluorescent SapC-DOPS Nanovesicles

We describe a multi-angle rotational optical imaging (MAROI) system for in vivo monitoring of physiopathological processes labeled with a fluorescent marker. Mouse models (brain tumor and arthritis) were used to evaluate the usefulness of this method. Saposin C (SapC)-dioleoylphosphatidylserine (DOPS) nanovesicles tagged with CellVue Maroon (CVM) fluorophore were administered intravenously. Animals were then placed in the rotational holder (MARS) of the in vivo imaging system. Images were acquired in 10&#176; steps over 380&#176;. A rectangular&#160;region of interest (ROI) was placed across the full image width at the model disease site. Within the ROI, and for every image, mean fluorescence intensity was computed after background subtraction. In the mouse models studied, the labeled nanovesicles were taken up in both the orthotopic and transgenic brain tumors, and in the arthritic sites (toes and ankles). Curve analysis of the multi angle image ROIs determined the angle with the highest signal. Thus, the optimal angle for imaging each disease site was characterized. The MAROI method applied to imaging of fluorescent compounds is a noninvasive, economical, and precise tool for in vivo quantitative analysis of the disease states in the described mouse models.

In Vivo Optical Imaging of Brain Tumors and Arthritis Using Fluorescent SapC-DOPS Nanovesicles

We describe a multi-angle rotational optical imaging (MAROI) system for in vivo quantitation of a fluorescent marker delivered by saposin C (SapC)-dioleoylphosphatidylserine (DOPS) nanovesicles. Employing mouse models of cancer and arthritis, we demonstrate how the MAROI signal curve analysis can be used for the precise mapping and biological characterization of disease processes.

We describe a multi-angle rotational optical imaging (MAROI) system for in vivo monitoring of physiopathological processes labeled with a fluorescent ...

Combined In vivo Optical and &#181;CT Imaging to Monitor Infection, Inflammation, and Bone Anatomy in an Orthopaedic Implant Infection in Mice

Multimodality imaging has emerged as a common technological approach used in both preclinical and clinical research. Advanced techniques that combine in vivo optical and &#956;CT imaging allow the visualization of biological phenomena in an anatomical context. These imaging modalities may be especially useful to study conditions that impact bone. In particular, orthopaedic implant infections are an important problem in clinical orthopaedic surgery. These infections are difficult to treat because bacterial biofilms form on the foreign surgically implanted materials, leading to persistent inflammation, osteomyelitis and eventual osteolysis of the bone surrounding the implant, which ultimately results in implant loosening and failure. Here, a mouse model of an infected orthopaedic prosthetic implant was used that involved the surgical placement of a Kirschner-wire implant into an intramedullary canal in the femur in such a way that the end of the implant extended into the knee joint. In this model, LysEGFP mice, a mouse strain that has EGFP-fluorescent neutrophils, were employed in conjunction with a bioluminescent Staphylococcus aureus strain, which naturally emits light. The bacteria were inoculated into the knee joints of the mice prior to closing the surgical site. In vivo bioluminescent and fluorescent imaging was used to quantify the bacterial burden and neutrophil inflammatory response, respectively. In addition, &#956;CT imaging was performed on the same mice so that the 3D location of the bioluminescent and fluorescent optical signals could be co-registered with the anatomical &#956;CT images. To quantify the changes in the bone over time, the outer bone volume of the distal femurs were measured at specific time points using a semi-automated contour based segmentation process. Taken together, the combination of in vivo bioluminescent/fluorescent imaging with &#956;CT imaging may be especially useful for the noninvasive monitoring of the infection, inflammatory response and anatomical changes in bone over time.

Combined In vivo Optical and  &amp;#181;CT Imaging to Monitor Infection, Inflammation, and Bone Anatomy in an Orthopaedic Implant Infection in Mice

Combined optical and &#956;CT imaging in a mouse model of orthopaedic implant infection, utilizing a bioluminescent engineered strain of Staphylococcus aureus, provided the capability to noninvasively and longitudinally monitor the dynamics of the bacterial infection, as well as the corresponding inflammatory response and anatomical changes in the bone.

Multimodality imaging has emerged as a common technological approach used in both preclinical and clinical research. Advanced techniques that combine in ...

Cerenkov Luminescence Imaging of Interscapular Brown Adipose Tissue

Brown adipose tissue (BAT), widely known as a &#8220;good fat&#8221; plays pivotal roles for thermogenesis in mammals. This special tissue is closely related to metabolism and energy expenditure, and its dysfunction is one important contributor for obesity and diabetes. Contrary to previous belief, recent PET/CT imaging studies indicated the BAT depots are still present in human adults. PET imaging clearly shows that BAT has considerably high uptake of 18F-FDG under certain conditions. In this video report, we demonstrate that Cerenkov luminescence imaging (CLI) with 18F-FDG can be used to optically image BAT in small animals. BAT activation is observed after intraperitoneal injection of norepinephrine (NE) and cold treatment, and depression of BAT is induced by long anesthesia. Using multiple-filter Cerenkov luminescence imaging, spectral unmixing and 3D imaging reconstruction are demonstrated. Our results suggest that CLI with 18F-FDG is a practical technique for imaging BAT in small animals, and this technique can be used as a cheap, fast, and alternative imaging tool for BAT research.

In this video report, we show the application of Cerenkov Luminescence Imaging (CLI) for interscapular brown adipose tissue in mice under activated and depressed conditions.

Brown adipose tissue (BAT), widely known as a &#8220;good fat&#8221; plays pivotal roles for thermogenesis in mammals. This special tissue is closely ...

Busulfan as a Myelosuppressive Agent for Generating Stable High-level Bone Marrow Chimerism in Mice

Bone marrow transplantation (BMT) is often used to replace the bone marrow (BM) compartment of recipient mice with BM cells expressing a distinct biomarker isolated from donor mice. This technique allows for identification of donor-derived hematopoietic cells within the recipient mice, and can be used to isolate and characterize donor cells using various biochemical techniques. BMT typically relies on myeloablative conditioning with total body irradiation to generate niche space within the BM compartment of recipient mice for donor cell engraftment. The protocol we describe here uses myelosuppressive conditioning with the chemotherapeutic agent busulfan. Unlike irradiation, which requires the use of specialized facilities, busulfan conditioning is performed using intraperitoneal injections of 20 mg/kg busulfan until a total dose of 60-100 mg/kg has been administered. Moreover, myeloablative irradiation can have toxic side effects and requires successful engraftment of donor cells for survival of recipient mice. In contrast, busulfan conditioning using these doses is generally well tolerated and mice survive without donor cell support. Donor BM cells are isolated from the femurs and tibiae of mice ubiquitously expressing green fluorescent protein (GFP), and injected into the lateral tail vein of conditioned recipient mice. BM chimerism is estimated by quantifying the number of GFP+ cells within the peripheral blood following BMT. Levels of chimerism&nbsp;>80% are typically observed in the peripheral blood 3-4 weeks post-transplant and remain established for at least 1 year. As with irradiation, conditioning with busulfan and BMT allows for the accumulation of donor BM-derived cells within the central nervous system (CNS), particularly in mouse models of neurodegeneration. This busulfan-mediated CNS accumulation may be more physiological than total body irradiation, as the busulfan treatment is less toxic and CNS inflammation appears to be less extensive. We hypothesize that these cells can be genetically engineered to deliver therapeutics to the CNS.

We describe a protocol whereby busulfan conditioning permits the bone marrow of a recipient mouse to be replaced with bone marrow cells from donor mice ubiquitously expressing green fluorescent protein, in the absence of irradiation. This technique is useful to study bone marrow cell accumulation in the central nervous system.

Bone marrow transplantation (BMT) is often used to replace the bone marrow (BM) compartment of recipient mice with BM cells expressing a distinct ...

Rose Bengal Photothrombosis by Confocal Optical Imaging In Vivo: A Model of Single Vessel Stroke

In vivo imaging techniques have increased in utilization due to recent advances in imaging dyes and optical technologies, allowing for the ability to image cellular events in an intact animal. Additionally, the ability to induce physiological disease states such as stroke in vivo increases its utility. The technique described herein allows for physiological assessment of cellular responses within the CNS following a stroke and can be adapted for other pathological conditions being studied. The technique presented uses laser excitation of the photosensitive dye Rose Bengal in vivo to induce a focal ischemic event in a single blood vessel. 
The video protocol demonstrates the preparation of a thin-skulled cranial window over the somatosensory cortex in a mouse for the induction of a Rose Bengal photothrombotic event keeping injury to the underlying dura matter and brain at a minimum. Surgical preparation is initially performed under a dissecting microscope with a custom-made surgical/imaging platform, which is then transferred to a confocal microscope equipped with an inverted objective adaptor. Representative images acquired utilizing this protocol are presented as well as time-lapse sequences of stroke induction. This technique is powerful in that the same area can be imaged repeatedly on subsequent days facilitating longitudinal in vivo studies of pathological processes following stroke.

Rose Bengal Photothrombosis by Confocal Optical Imaging In Vivo:  A Model of Single Vessel Stroke

Here, we describe a semi-invasive optical microscopy approach for the induction of a Rose Bengal photothrombotic clot in the somatosensory cortex of a mouse in vivo. The technical aspects of the imaging procedure are described from induction of a photothrombotic event to application and data collection.

In vivo imaging techniques have increased in utilization due to recent advances in imaging dyes and optical technologies, allowing for the ability to ...

A Neonatal Mouse Spinal Cord Compression Injury Model

Spinal cord injury (SCI) typically causes devastating neurological deficits, particularly through damage to fibers descending from the brain to the spinal cord. A major current area of research is focused on the mechanisms of adaptive plasticity that underlie spontaneous or induced functional recovery following SCI. Spontaneous functional recovery is reported to be greater early in life, raising interesting questions about how adaptive plasticity changes as the spinal cord develops. To facilitate investigation of this dynamic, we have developed a SCI model in the neonatal mouse. The model has relevance for pediatric SCI, which is too little studied. Because neural plasticity in the adult involves some of the same mechanisms as neural plasticity in early life1, this model may potentially have some relevance also for adult SCI. Here we describe the entire procedure for generating a reproducible spinal cord compression (SCC) injury in the neonatal mouse as early as postnatal (P) day 1. SCC is achieved by performing a laminectomy at a given spinal level (here described at thoracic levels 9-11) and then using a modified Yasargil aneurysm mini-clip to rapidly compress and decompress the spinal cord. As previously described, the injured neonatal mice can be tested for behavioral deficits or sacrificed for ex vivo physiological analysis of synaptic connectivity using electrophysiological and high-throughput optical recording techniques1. Earlier and ongoing studies using behavioral and physiological assessment have demonstrated a dramatic, acute impairment of hindlimb motility followed by a complete functional recovery within 2 weeks, and the first evidence of changes in functional circuitry at the level of identified descending synaptic connections1.

This article describes a method for generating a reproducible spinal cord compression injury (SCI) in the neonatal mouse. The model provides an advantageous platform for studying mechanisms of adaptive plasticity that underlie spontaneous functional recovery.

Spinal cord injury (SCI) typically causes devastating neurological deficits, particularly through damage to fibers descending from the brain to the spinal ...

Quantitative Visualization of Leukocyte Infiltrate in a Murine Model of Fulminant Myocarditis by Light Sheet Microscopy

Light-sheet fluorescence microscopy (LSFM), in combination with chemical clearing protocols, has become the gold standard for analyzing fluorescently labelled structures in large biological specimens, and is down to cellular resolution. Meanwhile, the constant refinement of underlying protocols and the enhanced availability of specialized commercial systems enable us to investigate the microstructure of whole mouse organs and even allow for the characterization of cellular behavior in various live-cell imaging approaches. Here, we describe a protocol for the spatial whole-mount visualization and quantification of the CD45+ leukocyte population in inflamed mouse hearts. The method employs a transgenic mouse strain (CD11c.DTR)that has recently been shown to serve as a robust, inducible model for the study of the development of fulminant fatal myocarditis, characterized by lethal cardiac arrhythmias. This protocol includes myocarditis induction, intravital antibody-mediated cell staining, organ preparation, and LSFM with subsequent computer-assisted image post-processing. Although presented as a highly-adapted method for our particular scientific question, the protocol represents the blueprint of an easily adjustable system that can also target completely different fluorescent structures in other organs and even in other species.

Here, we describe a light-sheet microscopy approach to visualize the cardiac CD45+ leukocyte infiltrate in a murine model of aseptic fulminant myocarditis, which is induced by the intratracheal diphtheria toxin treatment of CD11c.DTR mice.

Light-sheet fluorescence microscopy (LSFM), in combination with chemical clearing protocols, has become the gold standard for analyzing fluorescently ...

High-Throughput Analysis of Optical Mapping Data Using ElectroMap

Optical mapping is an established technique for high spatio-temporal resolution study of cardiac electrophysiology in multi-cellular preparations. Here we present, in a step-by-step guide, the use of ElectroMap for analysis, quantification, and mapping of high-resolution voltage and calcium datasets acquired by optical mapping. ElectroMap analysis options cover a wide variety of key electrophysiological parameters, and the graphical user interface allows straightforward modification of pre-processing and parameter definitions, making ElectroMap applicable to a wide range of experimental models. We show how built-in pacing frequency detection and signal segmentation allows high-throughput analysis of entire experimental recordings, acute responses, and single beat-to-beat variability. Additionally, ElectroMap incorporates automated multi-beat averaging to improve signal quality of noisy datasets, and here we demonstrate how this feature can help elucidate electrophysiological changes that might otherwise go undetected when using single beat analysis. Custom modules are included within the software for detailed investigation of conduction, single file analysis, and alternans, as demonstrated here. This software platform can be used to enable and accelerate the processing, analysis, and mapping of complex cardiac electrophysiology.

This protocol describes the setup and use of ElectroMap, a MATLAB-based open-source software platform for analysis of cardiac optical mapping data. ElectroMap provides a versatile high-throughput tool for analysis of optical mapping voltage and calcium datasets across a wide range of cardiac experimental models.

Optical mapping is an established technique for high spatio-temporal resolution study of cardiac electrophysiology in multi-cellular preparations. Here we ...