authors gratefully acknowledge funding support from the Molly Towell Perinatal Research Foundation, Canadian Institutes of Health Research (CIHR), and Fonds de Recherche du Qu&#233;bec &#8211; Sant&#233; (FRQS) (to MGF) and CIHR-Quebec Training Network in Perinatal Research (QTNPR) (to LDD).
The authors wish to thank Esther Simard, Marco Bosa, Carl Bernard and Carmen Movila for technical assistance.

Animal care followed the guidelines of the Canadian Council on Animal Care and the approval by the Universit&#233; de Montr&#233;al Council on Animal Care (protocol #10-Rech-1560). Detailed information on materials and methods used is provided in the Table 1.
1. Anesthesia
<ol>
	<li>Insert a single-lumen catheter into a jugular vein.</li>
	<li>Sedate the ewe using acepromazine (Atravet 10 mg/mL) 2 mg intravenously approximately 30 min&#160;prior to the induction of anesthesia in order to reduce stress associated with the procedure which in turn reduces levels of cortisol.</li>
	<li>Administer diazepam (Diazepam 5 mg/mL) 20 mg, ketamine (Ketalar 100 mg/mL)&#160;4-5 mg/kg and propofol (Propofol 10 mg/mL) 0.5 to 1 mg/kg intravenously to induce general anesthesia.</li>
	<li>Insert an airway exchange catheter into the trachea using a laryngoscope with a Wisconsin type blade (Extra-long 350 mm Left-Handed Blade) to aid with the intubation. Slide the silicon endotracheal tube (9 to 12 mm Inner Diameter) off the airway exchange catheter and into the trachea. This technique facilitates the intubation process. Inflate the cuff of the endotracheal tube carefully to avoid pressure-induced ulceration of the trachea and fixate the tube to the head of the ewe.</li>
	<li>Connect the endotracheal tube to the respiratory circuit of the anesthetic machine and begin mechanical ventilation immediately. Adjust ventilator settings to maintain a PaCO2 within normal limits of 35 to 45 mmHg.</li>
	<li>Insert a catheter into the auricular artery (22 to 20 G; 1 in [0.9 x 25 mm] to 1.16 in [1.1 x 30 mm]) and connect to non-compliant tubing to monitor direct arterial blood pressure.</li>
	<li>Use a multi-parameter physiologic monitor to record the electrocardiogram, direct arterial blood pressure, oxygen saturation (SpO2), capnography (PETCO2), and temperature every 5 min. Transfer all physiologic data via a serial cable to a central physiologic data-collecting computer. Maintain normal body temperature using a circulating water blanket.</li>
	<li>Administer a balanced poly-ionic solution at 10 mL/kg for the first hour of general anesthesia and then reduce to 5 mL/kg/h.</li>
	<li>Administer trimethoprim-sulfadoxine 5 mg/kg IV to the ewe just prior to the skin incision as antibiotic prophylaxis.</li>
	<li>Use standard aseptic techniques with all surgical manipulations of the ewe and fetus.</li>
	<li>Barrier nurse the ewes at all times. This includes also the non-surgical personnel. This will minimize the enzootic potential for e.g.,&#160;Coxiella burnetii. Use gloves and masks (N-95 type) at all times.</li>
</ol>
2. Overview of the Surgical Procedure
<ol>
	<li>Make a 20 cm midline incision through the lower abdominal wall immediately cranial to the udder through the linea alba to minimise abdominal muscle damage.
	<ol>
		<li>Retract the greater omentum cranially and the uterine horns are palpated manually. Palpate each horn from the body of the uterus to the tip of the horn noticing the number of fetus and their size. If there is more than one fetus, choose the larger fetus by evaluating manually the head size and the width between the orbits.</li>
		<li>Hold the head of the chosen fetus firmly through the uterus partially exteriorized. Perform a 10 cm hysterotomy on the large curvature with Metzenbaum scissors. Put immediately over the head a non-latex sterile surgical glove filled with sterile saline as if it were a hand. Alternatively, use moistened 4x4s to keep the fetal head moist. Secure the uterus to the abdominal wall.
		<ol>
			<li>Exteriorize the right and left thoracic limbs and pull the fetus gently out of the uterus up to the xyphoid process.</li>
		</ol>
		</li>
		<li>Insert polyvinyl catheters into the right and left brachial vein and arteries using a standard cut-down technique. Insert another polyvinyl catheter into the amniotic cavity by fixating its&#8217; end to the sternum of the fetus.
		<ol>
			<li>Use sterile catheters only. We recommend gas sterilization in your standard facility. Attach each catheter to a needle and the needle to a double stopcock to allow for later blood sampling or pressure monitoring.</li>
		</ol>
		</li>
		<li>Suture stainless steel electrodes to the manubrium, xiphoid process and to each point of the shoulder to monitor the electrocardiogram (ECG).</li>
		<li>Return the fetus to the uterus. All catheters and electrodes exit via a small stab incision in the left flank.</li>
		<li>Close the laparotomy incision using a three layer closure. Suture the abdominal wall with a synthetic absorbable monofilament suture USP 2 in a simple continuous fashion. Close the subcutaneous space with a synthetic absorbable braided suture USP 0 in a simple continuous fashion. Use surgical stainless steel staples to close the skin incision.</li>
	</ol>
	</li>
	<li>Detailed description of the surgical procedure
	<ol>
		<li>Remove wool by shaving the xyphoid process to the mammary gland and along the fold of the flank on each side with a blade # 40. Clean the ventral abdomen then thoroughly with a 4% chlorhexidine gluconate and a soft brush for 3 min.
		<ol>
			<li>Perform a standard sterile scrub with chlorhexidine gluconate 4% starting from the center of the abdomen and progressing in a centrifugal fashion for 3 min. Pour sterile saline on the abdomen to remove the disinfectant soap. For the final step of the surgical preparation, perform three alternate passages of Chlorhexidine gluconate solution 2% and isopropyl alcohol 70%.</li>
		</ol>
		</li>
		<li>Ensure the depth of anesthesia is adequate prior to incision. Make a standard laparotomy incision from the umbilicus to just cranial to the udder through the linea alba to minimise abdominal muscle damage.</li>
		<li>Insert a long sponge forceps in the abdomen along the left abdominal wall up to the planned exit site for catheters in the paracostal region.
		<ol>
			<li>Push the tips of the forceps against the wall until an assistant can locate it and confirm the appropriate site. Open slightly (1 cm) the jaws of the forceps and let the assistant make a 2 cm full-thickness stab incision.</li>
			<li>Exteriorize the tips of the forceps through the incision, open again, and gently grasp the catheters with the forceps that is finally pulled out of the abdomen through the ventral abdominal incision. Of note, some groups implant the catheters and then exteriorize them. This has the drawback that intrasurgical fetal ECG monitoring is not possible.</li>
		</ol>
		</li>
		<li>Palpate the uterus to determine fetal position and numbers. Determine the largest fetus using the inter-aural distance. Incise the uterine wall on the large curvature over the dorsum of the head, avoiding the cotyledons.
		<ol>
			<li>Insert a blunt-ended cannula through the placental membranes to obtain an amniotic fluid sample free from hemorrhage. Incise the placental membranes using scissors.</li>
		</ol>
		</li>
		<li>Exteriorize the cranial half of the fetus through this incision. Place a sterile non latex surgical glove filled with sterile saline at 37 &#176;C over the fetus head to help maintain normothermia.
		<ol>
			<li>During removal of the fetal upper body from the uterus, have the assistant hold the Babcock&#39;s up to prevent the loss of amniotic fluid. Then, again using Babcock forceps, clamp the fetal membranes and uterine wall to the skin to prevent abdominal contamination with amniotic fluid.</li>
			<li>Expose only the parts of the fetal body that need to be instrumented and keep the remaining parts inside the uterus or covered by moist and warm (37 &#176;C) sterile cloths, respectively.</li>
		</ol>
		</li>
		<li>Abduct both thoracic limbs to facilitate the exposure to the brachial artery and vein bilaterally. Incise along the medial aspect of both antebrachium and carefully dissect around the brachial artery and vein.</li>
		<li>Insert polyvinyl catheters into the right and left brachial arteries and the left brachial vein using a standard cut-down technique.
		<ol>
			<li>Free the vessel to be catheterized from adjacent tissue over 1 cm. Ligate the distal portion of the vessel with a braided synthetic absorbable USP 2-0 suture. Preplace a ligature at the proximal aspect of the vessel but keep it untied.
			<ol>
				<li>Using Castroviejo scissors, cut the vessel transversally to approximately 30% of its diameter. Stop the blood flow partially by pulling on the proximal suture. Insert the catheter in a proximal direction.</li>
			</ol>
			</li>
			<li>Insert the polyvinyl catheter up to 8 cm proximally or until resistance is detected and then pull back slightly. Temporally secure the proximal aspect of the catheter using a vascular clamp. Place another suture around both the proximal and distal aspect of the catheter. An assistant is continuously aspirating and flushing the catheter to ensure patency of the catheter.</li>
			<li>Close the fetal skin using a USP 2-0 braided synthetic absorbable suture, with a continuous suture pattern.</li>
		</ol>
		</li>
		<li>To the right and left shoulder, manubrium and xyphoid process, fixate an insulated stainless steel electrode to facilitate the monitoring of the fetal ECG.</li>
		<li>Suture the amniotic pressure and sampling catheter to the sternum. This catheter is fenestrated at its extremity.</li>
		<li>Secure all the catheters on the dorsum of the fetus using a USP 2-0 braided synthetic absorbable suture material.</li>
		<li>Prior to the replacement of the fetus back into the uterus, administer clenbuterol 30 &#181;g IV slowly over 15 min&#160;to avoid hypotension and to provide uterine relaxation.</li>
		<li>Suture the fetal membranes using USP 4-0 braided synthetic absorbable suture material with a continuous pattern. Incorporate only one catheter or electrode at a time into the closure to ensure a tight closure. Use a double layer Cushing pattern respecting Halsted principles to close the uterine muscular layer using a USP 0 braided synthetic absorbable suture material. Bury the surgical knots carefully.</li>
		<li>Using a purse-string suture pattern, secure all the catheters and ECG cables as they exit the left paracostal incision.</li>
		<li>Using a USP 2&#160;monofilament synthetic absorbable suture material secure the linea alba with a continuous pattern. Close the subcutaneous tissues using USP 2-0 braided synthetic absorbable suture material with a continuous pattern. Secure the skin layer with surgical staples.</li>
		<li>Administer 250 mg of ampicillin intravenously and again via the amniotic catheter into the amniotic cavity. Replace lost amniotic fluids with warm saline.</li>
		<li>Place all the exteriorized catheters and ECG electrodes into a place bag to maintain sterility. Place a stockinette around the torso of the ewe to secure all the catheters and electrodes to the body of the ewe.</li>
		<li>Stop general anesthesia, and extubate the ewe once laryngeal reflexes have returned to normal.</li>
		<li>Return the ewe to a metabolic cage once she is stable following general anesthesia. The ewe will reside in the metabolic cage for the duration of the experiment. The ewe should be able to stand, lie down and eat ad libitum while monitoring the non-anesthetized fetus without sedating the mother.</li>
		<li>For the following three days, administer antibiotics prophylactically to the ewe (Trimethoprim sulfadoxine 5 mg/kg) and fetus (250 mg of ampicillin intravenously and again via the amniotic catheter).</li>
		<li>Evaluate the metabolic status of both ewe and fetus using blood gas analyses.</li>
		<li>Flush all catheters with the minimal volume of heparinized saline possible. Caution &#8211; do not exceed the daily dose of heparin and fluids permissible to the fetus. It is possible to fluid overload the fetus. Flush slowly once a day ~5 mL NaCl per line after antibiotic prophylaxis.</li>
	</ol>
	</li>
	<li>Data recording and analysis
	<ol>
		<li>During surgery, optionally record the maternal and fetal ECG and heart rates as well as maternal arterial blood pressure and airway pressure (Paw) continuously (Figure 1). Use the Life Window Monitor to acquire all maternal data except ECG. Feed these data into analog-digital converter along with fetal and maternal ECG signals; pass&#160;maternal and fetal ECG first into 1901 pre-amplifier. Record and display all data in manufacturer&#8217;s software.</li>
		<li>Take a 1 mL arterial sample simultaneously from the ewe and fetus for arterial blood gas analysis, lactate, glucose and base excess determination (in plasma) at the begin of the fetal surgery (immediately after inserting the first arterial catheter) and after closing the uterus.</li>
		<li>During postoperative recovery, take a 3 mL fetal blood sample to measure the IL-6 and TNF-&#945; inflammatory profiles. Centrifuge the plasma at 4 &#176;C (4 min, 4,000 x g), decant and store the plasma at -80 &#176;C for subsequent ELISA testing. 
		Note: For the purpose of the reported representative results, six days after surgery the animals were sacrificed using intravenous injection of 20 mL sodium pentobarbital. Fetal growth was assessed by body, brain, liver and maternal weights. The duration of the experimental period will obviously vary depending on the design chosen for the particular research question and can reach ~6 weeks.</li>
	</ol>
	</li>
	<li>Cytokine analyses (optional step)
	<ol>
		<li>Determine cytokine concentrations (IL-6, TNF-&#945;) in plasma using an ovine-specific sandwich&#160;ELISA. Pre-coat mouse anti-sheep monoclonal antibodies (capture antibody IL-6) or mouse anti-bovine monoclonal antibody (TNF-&#945;) at a concentration 4 &#181;g/ml on ELISA plate at 4 &#176;C for O/N, after 3 times wash with washing buffer (0.05% Tween 20 in PBS, PBST).</li>
		<li>Block the plates for 1 h with 1% BSA in PBST. Wash the plates with washing buffer 3 times.</li>
		<li>Use recombinant sheep proteins (IL-6, TNF-&#945;) as ELISA standard, prepare a serial dilutions range from standard 1 of 2,000 ng/ml to standard 7 of 31.25 pg/ml.</li>
		<li>Load 50 &#181;l of serial diluted protein standards and samples per well and incubate for 2 hr at RT, wash the plates 3 times. Run all standards and samples in duplicate.</li>
		<li>Apply 50 &#181;l of rabbit anti-sheep polyclonal antibodies (detection antibody IL-6) or rabbit anti-bovine polyclonal antibody (TNF-&#945;) at a dilution of 1:250 in wells and incubate for 30 min at RT. Wash the plates with washing buffer 5 times.</li>
		<li>Add 50 &#181;l of the goat anti-rabbit IgG-HRP conjugated (dilution 1:5,000) for 30 min.</li>
		<li>Incubate with 50 &#181;l of TMB substrate solution per well.</li>
		<li>Stop colour development reaction at desired time with 25 &#181;l of 2 N sulphuric acid.</li>
		<li>Read the plates on ELISA plate reader at 450 nm, with a 570 nm wavelength correction. 
		Note: In our assays, the sensitivity of IL-6 ELISA was 16 pg/ml, the sensitivity of TNF-&#945; ELISA was 13.9 pg/ml, respectively. For all assays, the intra-assay and inter-assay coefficients of variance was &#60;5% and &#60;10%, respectively.</li>
	</ol>
	</li>
	<li>Statistical analyses 
	Note: The exact statistical methods will depend on the research question. Here we report the methods used to test for any significant differences in the blood gases reported in Table 2.
	<ol>
		<li>Test normal data distribution using Kolmogorov-Smirnov test followed by parametric or non-parametric tests with adjustment for multiple comparisons, as appropriate.</li>
		<li>Use K-means cluster analysis to identify the fetuses in the cohort that were spontaneously hypoxic and determine the respective pO2 and O2Sat values (consider pO2 &#60; 20 mmHg or O2Sat &#60; 55% as fetal hypoxia). 44-48</li>
		<li>Use statistical software such as SPSS for the analyses.</li>
		<li>Present data as Mean&#177;SD with statistical differences at p&#60;0.05 considered significant.</li>
	</ol>
	</li>
</ol>

No disclosures have been made.

The anesthetic and surgical procedures are presented that are required for establishing an animal model for studying fetal physiology and pathophysiology: the chronically instrumented non-anesthetized fetal sheep.
Four critical steps within the protocol should be emphasized. First, passing the catheters and electrodes through the maternal flank: it is important that this is done at once to avoid any internal organ injuries. Second, securing the uterotomy operating site prior to exteriorizing t...

A variety of animal models exist for the study of both normal and compromised pregnancies, including laboratory rodents, non-human primates and domestic ruminants.1,2,3,4,5 The chronically instrumented pregnant sheep has been used extensively for 50 years as a model of human fetal development and responses to pathophysiologic stimuli such as lipopolysaccharide (LPS).6-10 The lesions following LPS exposure mimic exactly what is seen in preterm infants with periventricular leukomalacia, which is due to a similar maturational profile of both species.11, 12
Other pregnancy complications have also been studied in great detail such as the discovery that antenatal glucocorticoids promote lung development 13-15 and understanding the impact of intrauterine growth restriction (IUGR) on the fetus 16,17.
The extensive use of the fetal sheep model is due to the unique amenability of the non-anesthetized fetal sheep to the surgical placement and maintenance of catheters and electrodes, allowing repetitive blood sampling, recording of bioelectrical activity, application of electric stimulation and in vivo brain imaging.18 Telemetry is also possible, although less frequently used yet due to the higher sophistication to set up as well as the initial and maintenance cost.19
Moreover, the fetal sheep model is very versatile as many variations of instrumentation are possible depending on the measures of interest. For example, it is possible to record over days to weeks multivariate signals in real time such as fetal breathing movements, electrical brain activity, cardiovascular responses, electrocardiogram, regional blood flow to a range of organs using flow probes or microspheres, etc. Thanks to this versatility, a wide range of studies have been conducted including the development of the cardiovascular system20,21, hypothalamo-pituitary-adrenal (HPA) axis22, brain development23 and sleep states development in particular24, effects of hypoxia/asphyxia25, therapeutic hypothermia 26, inflammation6-11, combination of both27, glucocorticoids28,29, anti-depressants30, broncho-pulmonary dysplasia (BPD) 31,32, fetal programming33,34,35,36,37,38,39 or development of novel fetal monitoring modalities prior and during labor to name but a few areas of investigation. 40,41,42,43
The overall goal of the method presented is to show this versatile model&#8217;s basic implementation. It allows establishing a wide variety of acute and chronic experimental protocols studying fetal physiology and pathophysiology on the integrative, organ, cellular and molecular levels.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>ACE Light source</td>
			<td>Schott-Fostec</td>
			<td>A20500</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting scissors</td>
			<td>Fine Science Tools</td>
			<td>14060 - 11</td>
			<td></td>
		</tr>
		<tr>
			<td>Angled dissecting scissors</td>
			<td>Fine Science Tools</td>
			<td>15006 - 09</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel handle</td>
			<td>Fine Science Tools</td>
			<td>10003 - 12</td>
			<td>alternating dissecting tool</td>
		</tr>
		<tr>
			<td>Curved scalpel blades #12</td>
			<td>Fine Science Tools</td>
			<td>10012 - 00</td>
			<td>alternating dissecting tool</td>
		</tr>
		<tr>
			<td>Bone scissors</td>
			<td>Fine Science Tools</td>
			<td>16044 - 10</td>
			<td></td>
		</tr>
		<tr>
			<td>S &#38; T suture tying forceps</td>
			<td>Fine Science Tools</td>
			<td>00272 - 13</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont SS forceps - angled</td>
			<td>Fine Science Tools</td>
			<td>11203 - 25&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Braided silk suture size 6-0</td>
			<td>Teleflex Medical</td>
			<td>07 - 30&#160; - 10</td>
			<td></td>
		</tr>
		<tr>
			<td>Medical Tape</td>
			<td>transpore</td>
			<td>3M</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine hydrochloride 100 mg/ml</td>
			<td>Hospira</td>
			<td>NDC 0409 - 2051 - 05</td>
			<td>Final Does is 80 mg/kg</td>
		</tr>
		<tr>
			<td>Tranqui Ved Injection (xylazine 100 mg/ml)</td>
			<td>Vecdo</td>
			<td>NDC 50989 - 234 - 11</td>
			<td>Final Does is 10 mg/kg</td>
		</tr>
		<tr>
			<td>Reactive orange 14</td>
			<td>Sigma - Aldrich</td>
			<td>R - 8254</td>
			<td></td>
		</tr>
		<tr>
			<td>Ringers Solution Components</td>
			<td></td>
			<td></td>
			<td>Solution is gas equilibrated with 95% O2 and 5% Co2, final pH 7.4</td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Sigma - Aldrich</td>
			<td>S7653</td>
			<td>Final Concentration: 118 mM</td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Fisher Scientific</td>
			<td>P217 - 3</td>
			<td>Final Concentration: 4.7 mM</td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate</td>
			<td>Fisher Scientific</td>
			<td>C79 - 500</td>
			<td>Final Concentration: 2.5 mM</td>
		</tr>
		<tr>
			<td>Potassium phosphate monobasic</td>
			<td>Fisher Scientific</td>
			<td>P -285</td>
			<td>Final Concentration: 1.2 mM</td>
		</tr>
		<tr>
			<td>Magnesium sulfate</td>
			<td>J.T. Baker</td>
			<td>Jan-00</td>
			<td>Final Concentration: 0.57 mM</td>
		</tr>
		<tr>
			<td>4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES)</td>
			<td>Fisher Scientific</td>
			<td>BP 310 - 500</td>
			<td>Final Concentration: 5.95 g/L</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma - Aldrich</td>
			<td>G8270</td>
			<td>Final Concentration: 5.5 mM</td>
		</tr>
		<tr>
			<td>LifeWindow</td>
			<td>Digicare Biomedical Technology</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CED bioamplifier and ADC units</td>
			<td>Cambridge Electronic Design Limited, 
			Unit 4, Science Park, 
			Milton Road, 
			Cambridge CB4 0FE 
			ENGLAND.</td>
			<td>Bioamp: 1902; ADC: micro1401; Data acquisition software: Spike 2, V7.13</td>
			<td></td>
		</tr>
		<tr>
			<td>Neurolog analog signal bioamplifier</td>
			<td>Digitimer Ltd 
			37 Hydeway 
			Welwyn Garden City 
			Hertfordshire, AL7 3BE, England</td>
			<td>NL108A</td>
			<td></td>
		</tr>
		<tr>
			<td>ABL800Flex</td>
			<td>Radiometer Canada; 200 Aberdeen Dr, London, ON N5V 4N2</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf 5804R</td>
			<td>Eppendorf Canada; 2810 Argentia Road, #2 
			Mississauga, Ontario, L5N 8L2</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Arrow Jugular Catheterization Set</td>
			<td>Arrow International, Inc., 2400 Bernville Road, Reading, PA 19605 USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Atravet</td>
			<td></td>
			<td></td>
			<td>10 mg/mL</td>
		</tr>
		<tr>
			<td>Diazepam</td>
			<td></td>
			<td></td>
			<td>5mg/mL</td>
		</tr>
		<tr>
			<td>Ketamine</td>
			<td>Ketalar</td>
			<td></td>
			<td>100 mg/mL</td>
		</tr>
		<tr>
			<td>Propofol</td>
			<td></td>
			<td></td>
			<td>10 mg/mL</td>
		</tr>
		<tr>
			<td>SurgiVeT</td>
			<td>Endotracheal Tubes; Smiths Medical ASD, Inc. St. Paul, MN 55112, USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cook Airway Exchange Catheter with RAPI-FIT Adapters</td>
			<td>Cook Critical Care 750, Bloomington IN 47402-0489 USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dispomed Ventilator</td>
			<td>Dispomed Ltd., 745 Nazaire-Laurin, Joliette, Quebec J6E 0L6</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>BD Insyte-W</td>
			<td>Becton Dickinson, Infusion Therapy Systems Inc., 9450 S State St, Sandy Utah 84070 USA</td>
			<td></td>
			<td>22 to 20 G; 1 in [0.9 x 25 mm] to 1.16 in [1.1 x 30 mm]</td>
		</tr>
		<tr>
			<td>Edwards Lifesciences Ref: PX272 Pressure monitoring kit with TruWave Disposable Pressure</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>LifeWindow LW6000</td>
			<td>Digicare Biomedical Technology 107 Commerce Road, Boynton Beach, FL 33426-9365 USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Gaymar</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Babcock</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Polyvinyl catheters</td>
			<td>SCI (Scientific Commodities Inc.)</td>
			<td></td>
			<td>2 meters</td>
		</tr>
		<tr>
			<td>2-0 Vicryl</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Castroviejo scissors</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>electrocardiogram (ECG)</td>
			<td>LIFYY, Metrofunk Kabel-Union, Berlin, Germany</td>
			<td></td>
			<td>four copper electrodes in single sheath, 2 meters</td>
		</tr>
		<tr>
			<td>2-O Vicryl</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>3-0 Vicryl</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PDS II USP</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Trimethoprim sulfadoxine</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ampicillin</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Stopcock</td>
			<td>Argon Medical, Cat&#160;041220001A</td>
			<td></td>
			<td>Double 4-way Stopcock with male luer lock</td>
		</tr>
		<tr>
			<td>Needles</td>
			<td>Tyco Healthcare 8881202389</td>
			<td></td>
			<td>Monoject aluminum hub blunt needles, 22Gx, 0.7mmx 38.1mm: for fetal arterial and venous catheters</td>
		</tr>
		<tr>
			<td>Needles</td>
			<td>Tyco Healthcare 8881202322</td>
			<td></td>
			<td>Monoject aluminum hub blunt needles, 16Gx, 1.6mmx38.1mm: for amniotic catheters</td>
		</tr>
	</tbody>
</table>

instrumentation of near-term fetal sheep for multivariate chronic non-anesthetized recordings

The chronically instrumented pregnant sheep has been used as a model of human fetal development and responses to pathophysiologic stimuli such as endotoxins, bacteria, umbilical cord occlusions, hypoxia and various pharmacological treatments. The life-saving clinical practices of glucocorticoid treatment in fetuses at risk of premature birth and the therapeutic hypothermia have been developed in this model. This is due to the unique amenability of the non-anesthetized fetal sheep to the surgical placement and maintenance of catheters and electrodes, allowing repetitive blood sampling, substance injection, recording of bioelectrical activity, application of electric stimulation and in vivo organ imaging. Here we describe the surgical instrumentation procedure required to achieve a stable chronically instrumented non-anesthetized fetal sheep model including characterization of the post-operative recovery from blood gas, metabolic and inflammation standpoints.

38 pregnant time-dated ewes were instrumented at 128&#177;2 days of gestation (dGA, ~0.88 gestation, term 145 dGA) with arterial, venous and amniotic catheters and electrocardiogram (ECG) electrodes with sterile technique under general anesthesia (both ewe and fetus). In case of twin pregnancy the larger fetus was chosen based on palpating and estimating the intertemporal diameter; alternatively, the fetus to be instrumented may be selected randomly to avoid any potential bias or both fetuses may be instrumented. The tot...

Watch this Scientific Journal Video about Instrumentation of Near-term Fetal Sheep for Multivariate Chronic Non-anesthetized Recordings at JoVE.com

Instrumentation of Near-term Fetal Sheep for Multivariate Chronic Non-anesthetized Recordings

The chronically instrumented non-anesthetized fetal sheep model is used to study human fetal development in health and disease, because it permits surgical placement and maintenance of catheters and electrodes, repetitive blood sampling, substance injection, recording of bioelectrical activity, and in vivo imaging. We describe the procedures required to establish this model.

The chronically instrumented pregnant sheep has been used as a model of human fetal development and responses to pathophysiologic stimuli such as ...

instrumentation-near-term-fetal-sheep-for-multivariate-chronic-non

Research

JoVE Journal

Developmental Biology

9.4K Views.  CHUV, Université de Montréal, St-Hyacinthe, QC. The chronically instrumented non-anesthetized fetal sheep model is used to study human fetal development in health and disease, because it permits surgical placement and maintenance of catheters and electrodes, repetitive blood sampling, substance injection, recording of bioelectrical activity, and in vivo imaging. We describe the procedures required to establish this model.

Video: Instrumentation of Near-term Fetal Sheep for Multivariate Chronic Non-anesthetized Recordings

Preparation and Culture of Myogenic Precursor Cells/Primary Myoblasts from Skeletal Muscle of Adult and Aged Humans

Skeletal muscle homeostasis depends on muscle growth (hypertrophy), atrophy and regeneration. During ageing and in several diseases, muscle wasting occurs. Loss of muscle mass and function is associated with muscle fiber type atrophy, fiber type switching, defective muscle regeneration associated with dysfunction of satellite cells, muscle stem cells, and other pathophysiological processes. These changes are associated with changes in intracellular as well as local and systemic niches. In addition to most commonly used rodent models of muscle ageing, there is a need to study muscle homeostasis and wasting using human models, which due to ethical implications, consist predominantly of in vitro cultures. Despite the wide use of human Myogenic Progenitor Cells (MPCs) and primary myoblasts in myogenesis, there is limited data on using human primary myoblast and myotube cultures to study molecular mechanisms regulating different aspects of age-associated muscle wasting, aiding in the validation of mechanisms of ageing proposed in rodent muscle. The use of human MPCs, primary myoblasts and myotubes isolated from adult and aged people, provides a physiologically relevant model of molecular mechanisms of processes associated with muscle growth, atrophy and regeneration. Here we describe in detail a robust, inexpensive, reproducible and efficient protocol for the isolation and maintenance of human MPCs&#160;and their progeny&#160;&#8212; myoblasts and myotubes from human muscle samples using enzymatic digestion. Furthermore, we have determined the passage number at which primary myoblasts from adult and aged people undergo senescence in an in vitro culture. Finally, we show the ability to transfect these myoblasts and the ability to characterize their proliferative and differentiation capacity and propose their suitability for performing functional studies of molecular mechanisms of myogenesis and muscle wasting in vitro.

This protocol describes a robust, reproducible and simple method of isolation and culture of myoblast progenitor cells from the skeletal muscle of adult and aged people. The muscles used here include foot and leg muscles. This approach enables the isolation of an enriched population of primary myoblasts for functional studies.

Skeletal muscle homeostasis depends on muscle growth (hypertrophy), atrophy and regeneration. During ageing and in several diseases, muscle wasting ...

Isolation and Characterization of Mesenchymal Stromal Cells from Human Umbilical Cord and Fetal Placenta

The human umbilical cord (UC) and placenta are non-invasive, primitive and abundant sources of mesenchymal stromal cells (MSCs) that have increasingly gained attention because they do not pose any ethical or moral concerns. Current methods to isolate MSCs from UC yield low amounts of cells with variable proliferation potentials. Since UC is an anatomically-complex organ, differences in MSC properties may be due to the differences in the anatomical regions of their isolation. In this study, we first dissected the cord/placenta samples into three discrete anatomical regions: UC, cord-placenta junction (CPJ), and fetal placenta (FP). Second, two distinct zones, cord lining (CL) and Wharton&#39;s jelly (WJ), were separated. The explant culture technique was then used to isolate cells from the four sources. The time required for the primary culture of cells from the explants varied depending on the source of the tissue. Outgrowth of the cells occurred within 3 - 4 days of the CPJ explants, whereas growth was observed after 7 - 10 days and 11 - 14 days from CL/WJ and FP explants, respectively. The isolated cells were adherent to plastic and displayed fibroblastoid morphology and surface markers, such as CD29, CD44, CD73, CD90, and CD105, similarly to bone marrow (BM)-derived MSCs. However, the colony-forming efficiency of the cells varied, with CPJ-MSCs and WJ-MSCs showing higher efficiency than BM-MSCs. MSCs from all four sources differentiated into adipogenic, chondrogenic, and osteogenic lineages, indicating that they were multipotent. CPJ-MSCs differentiated more efficiently in comparison to other MSC sources. These results suggest that the CPJ is the most potent anatomical region and yields a higher number of cells, with greater proliferation and self-renewal capacities in vitro. In conclusion, the comparative analysis of the MSCs from the four sources indicated that CPJ is a more promising source of MSCs for cell therapy, regenerative medicine, and tissue engineering.

Here, we present a protocol for the dissection of human umbilical cord (UC) and fetal placenta sample into cord lining (CL), Wharton&#39;s jelly (WJ), cord-placenta junction (CPJ), and fetal placenta (FP) for the isolation and characterization of mesenchymal stromal cells (MSCs) using the explant culture technique.

The human umbilical cord (UC) and placenta are non-invasive, primitive and abundant sources of mesenchymal stromal cells (MSCs) that have increasingly ...

Whole-cell Patch-clamp Recordings of Isolated Primary Epithelial Cells from the Epididymis

The epididymis is an essential organ for sperm maturation and reproductive health. The epididymal epithelium consists of intricately connected cell types that are distinct not only in molecular and morphological features but also in physiological properties. These differences reflect their diverse functions, which together establish the necessary microenvironment for the post-testicular sperm development in the epididymal lumen. The understanding of the biophysical properties of the epididymal epithelial cells is critical for revealing their functions in sperm and reproductive health, under both physiological and pathophysiological conditions. While their functional properties have yet to be fully elucidated, the epididymal epithelial cells can be studied using the patch-clamp technique, a tool for measuring the cellular events and the membrane properties of single cells. Here, we describe the methods of cell isolation and whole-cell patch-clamp recording to measure the electrical properties of primary dissociated epithelial cells from the rat cauda epididymides.

We present a protocol that combines cell isolation and whole-cell patch-clamp recording to measure the electrical properties of the primary dissociated epithelial cells from the rat cauda epididymides. This protocol allows for investigation of the functional properties of primary epididymal epithelial cells to further elucidate the physiological role of the epididymis.

The epididymis is an essential organ for sperm maturation and reproductive health. The epididymal epithelium consists of intricately connected cell types ...

Zika Virus Infection of Cultured Human Fetal Brain Neural Stem Cells for Immunocytochemical Analysis

Human fetal brain neural stem cells are a unique non-genetically modified model system to study the impact of various stimuli on human developmental neurobiology. Rather than use an animal model or genetically modified induced pluripotent cells, human neural stem cells provide an effective in vitro system to examine the effects of treatments, screen drugs, or examine individual differences. Here, we provide the detailed protocols for methods used to expand human fetal brain neural stem cells in culture with serum-free media, to differentiate them into various neuronal subtypes and astrocytes via different priming procedures, and to freeze and recover these cells. Furthermore, we describe a procedure of using human fetal brain neural stem cells to study Zika virus infection.

This article details the methods that are used to expand human fetal brain neural stem cells in culture, as well as how to differentiate them into various neuronal subtypes and astrocytes, with an emphasis on the use of neural stem cells to study Zika virus infection.

Human fetal brain neural stem cells are a unique non-genetically modified model system to study the impact of various stimuli on human developmental ...

Intravenous and Intra-amniotic In Utero Transplantation in the Murine Model

In utero transplantation (IUT) is a unique and versatile mode of therapy that can be used to introduce stem cells, viral vectors, or any other substances early in the gestation. The rationale behind IUT for therapeutic purposes is based on the small size of the fetus, the fetal immunologic immaturity, the accessibility and proliferative nature of the fetal stem or progenitor cells, and the potential to treat a disease or the onset of symptoms prior to birth. Taking advantage of these normal developmental properties of the fetus, the delivery of hematopoietic stem cells (HSC) via an IUT has the potential to treat congenital hematologic disorders such as sickle cell disease, without the required myeloablative or immunosuppressive conditioning required for postnatal HSC transplants. Similarly, the accessibility of progenitor cells in multiple organs during development potentially allows for a more efficient targeting of stem/progenitor cells following an IUT of viral vectors for gene therapy or genome editing. Additionally, IUT can be used to study normal developmental processes including, but not limited to, the development of immunologic tolerance. The murine model provides a valuable and affordable means to understanding the potential and limitations of IUT prior to pre-clinical large animal studies and an eventual clinical application. Here, we describe a protocol for performing an IUT in the murine fetus through intravenous and intra-amniotic routes. This protocol has been used successfully to elucidate the necessary conditions and mechanisms behind in utero hematopoietic stem cell transplantation, tolerance induction, and in utero gene therapy.

Intravenous and Intra-amniotic In Utero Transplantation in the Murine Model

We describe a protocol for performing an in utero transplantation (IUT) through intravenous and intra-amniotic routes of injection in the murine model. This protocol can be used to introduce cells, viral vectors, and other substances into the unique immune-tolerant fetal environment.

In utero transplantation (IUT) is a unique and versatile mode of therapy that can be used to introduce stem cells, viral vectors, or any other substances ...

High Frequency Ultrasound for the Analysis of Fetal and Placental Development In Vivo

Ultrasound imaging is a widespread method used to detect organ anomalies and tumors in human and animal tissues. The method is non-invasive, harmless, and painless, and the application is easy, fast, and can be done anywhere, even with mobile devices. During pregnancy, ultrasound imaging is standardly used to closely monitor fetal development. The technique is important to assess intrauterine growth restriction (IUGR), a pregnancy complication with short- and long-term health consequences for both the mother and fetus. Understanding the process of IUGR is indispensable for developing effective therapeutic strategies. 
The ultrasound system used in this manuscript is an ultrasound device produced for the analysis of small animals and can be used in various research fields, including pregnancy research. Here we describe the usage of the system for in vivo analysis of fetuses from natural killer (NK) cell/mast cell (MC)-deficient mothers that give birth to growth-restricted pups. The protocol includes preparation of the system, handling of the mice before and during measurements, and the usage of the B-mode, color doppler mode, and pulse-wave doppler mode. Fetal size, placental size, and blood supply to the fetus were analyzed. We found reduced implantation sizes and smaller placentas in NK/MC-deficient mice from mid-gestation onwards. In addition, MC/NK-deficiency was associated with absent and reversed end diastolic flow in the fetal Arteria umbilicalis(UmA) and an elevated resistance index. The methods described in the protocol can easily be used for related and non-related research topics.

High Frequency Ultrasound for the Analysis of Fetal and Placental Development In Vivo

Here we describe the technique of high frequency ultrasound for in vivo analysis of fetuses in mice. This method allows the follow-up of fetuses and the analysis of placental parameters as well as maternal and fetal blood flow throughout pregnancy.

Ultrasound imaging is a widespread method used to detect organ anomalies and tumors in human and animal tissues. The method is non-invasive, harmless, and ...

Culturing and Measuring Fetal and Newborn Murine Long Bones

Long bones are complex and dynamic structures, which arise from endochondral ossification via a cartilage intermediate. The limited access to healthy human bones makes particularly valuable the use of mammalian models, such as mouse and rat, to look into different aspects of bone growth and homeostasis. Additionally, the development of sophisticated genetic tools in mice allows more complex studies of long bone growth and asks for an expansion of techniques used to study bone growth. Here, we present a detailed protocol for ex vivo murine bone culture, which allows the study of bone and cartilage in a tightly controlled manner while recapitulating most of the in vivo process. The method described allows the culture of a range of bones, including tibia, femur, and metatarsal bones, but we have focused mainly on tibial culture here. Moreover, it can be used in combination with other techniques, such as time-lapse live imaging or drug treatment.

Here, we present a method for ex vivo culture of long murine bones at both fetal and newborn stages, suitable for analyzing bone and cartilage development and homeostasis in controlled conditions while recapitulating the in vivo process.

Long bones are complex and dynamic structures, which arise from endochondral ossification via a cartilage intermediate. The limited access to healthy ...

Recapitulating Suckling-to-Weaning Transition In Vitro using Fetal Intestinal Organoids

At the end of the suckling period, many mammalian species undergo major changes in the intestinal epithelium that are associated with the capability to digest solid food. This process is termed suckling-to-weaning transition and results in the replacement of neonatal epithelium with adult epithelium which goes hand in hand with metabolic and morphological adjustments. These complex developmental changes are the result of a genetic program that is intrinsic to the intestinal epithelial cells but can, to some extent, be modulated by extrinsic factors. Prolonged culture of mouse primary intestinal epithelial cells from late fetal period, recapitulates suckling-to-weaning transition in vitro. Here, we describe a detailed protocol for mouse fetal intestinal organoid culture best suited to model this process in vitro. We describe several useful assays designed to monitor the change of intestinal functions associated with suckling-to-weaning transition over time. Additionally, we include an example of an extrinsic factor that is capable to affect suckling-to-weaning transition in vivo, as a representation of modulating the timing of suckling-to-weaning transition in vitro. This in vitro approach can be used to study molecular mechanisms of the suckling-to-weaning transition as well as modulators of this process. Importantly, with respect to animal ethics in research, replacing in vivo models by this in vitro model contributes to refinement of animal experiments and possibly to a reduction in the use of animals to study gut maturation processes.

This protocol describes how to mimic suckling-to-weaning transition in vitro using mouse late fetal intestinal organoids cultured for 30 days.

At the end of the suckling period, many mammalian species undergo major changes in the intestinal epithelium that are associated with the capability to ...

Transuterine Fetal Tracheal Occlusion Model in Mice

Fetal tracheal occlusion (TO), an established treatment modality, promotes fetal lung growth and survival in severe congenital diaphragmatic hernia (CDH). Following TO, retention of the secreted epithelial fluid increases luminal pressure and induces lung growth. Various animal models have been defined to understand the pathophysiology of CDH and TO. All have their own advantages and disadvantages such as the difficulty of the technique, the size of the animal, cost, high mortality rates, and the availability of genetic tools. Herein, a novel transuterine model of murine fetal TO is described. Pregnant mice were anesthetized, and the uterus exposed via a midline laparotomy. The trachea of selected fetuses were ligated with a single transuterine suture placed behind the trachea, one carotid artery, and one jugular vein. The dam was closed and allowed to recover. Fetuses were collected just before parturition. Lung to body weight ratio in TO fetuses was higher than that in control fetuses. This model provides researchers with a new tool to study the impact of both TO and increased luminal pressure on lung development.

Various animal models of congenital diaphragmatic hernia and fetal tracheal occlusion present advantages and disadvantages regarding ethical issues, cost, surgical difficulty, size, survival rates, and availability of genetic tools. This model provides a new tool to study the impact of both tracheal occlusion and increased luminal pressure on lung development.

Fetal tracheal occlusion (TO), an established treatment modality, promotes fetal lung growth and survival in severe congenital diaphragmatic hernia (CDH). ...

Vitrification of In Vitro Matured Oocytes Collected from Adult and Prepubertal Ovaries in Sheep

In livestock, in vitro embryo production systems can be developed and sustained thanks to the large number of ovaries and oocytes that can be easily obtained from a slaughterhouse. Adult ovaries always bear several antral follicles, while in pre-pubertal donors the maximal numbers of oocytes are available at 4 weeks of age, when ovaries bear peak numbers of antral follicles. Thus, 4 weeks old lambs are considered good donors, even if the developmental competence of prepubertal oocytes is lower compared to their adult counterpart. 
Basic research and commercial applications would be boosted by the possibility of successfully cryopreserving vitrified oocytes obtained from both adult and prepubertal donors. The vitrification of oocyte collected from prepubertal donors would also allow shortening the generation interval and thus increasing the genetic gain in breeding programs. However, the loss of developmental potential after cryopreservation makes mammalian oocytes probably one of the most difficult cell types to cryopreserve. Among the available cryopreservation techniques, vitrification is widely applied to animal and human oocytes. Despite recent advancements in the technique, exposures to high concentrations of cryoprotective agents as well as chilling injury and osmotic stress still induce several structural and molecular alterations and reduce the developmental potential of mammalian oocytes. Here, we describe a protocol for the vitrification of sheep oocytes collected from juvenile and adult donors and matured in vitro prior to cryopreservation. The protocol includes all the procedures from oocyte in vitro maturation to vitrification, warming and post-warming incubation period. Oocytes vitrified at the MII stage can indeed be fertilized following warming, but they need extra time prior to fertilization to restore damage due to cryopreservation procedures and to increase their developmental potential. Thus, post-warming culture conditions and timing are crucial steps for the restoration of oocyte developmental potential, especially when oocyte are collected from juvenile donors.

The protocol aims at providing a standard method for the vitrification of adult and juvenile sheep oocytes. It includes all the steps from the preparation of the in vitro maturation media to the post-warming culture. Oocytes are vitrified at the MII stage using Cryotop to ensure the minimum essential volume.

In livestock, in vitro embryo production systems can be developed and sustained thanks to the large number of ovaries and oocytes that can be easily ...