What surgical instrument is used to hold delicate tissue such as the intestine?

Scissors

Surgical scissors are required for cutting skin, undermining the subcutis and deeper fascial layers, cutting sutures, and removing wound dressings (Fig. 144.12). Scissors may have long or short handles, and the blades are straight or curved and serrated or smooth. The tips may be sharp or blunt. Scissors used in cutaneous surgery can be either completely stainless steel (most popular, least expensive) or have tungsten carbide inserts to strengthen the blades.

Gradle scissors are small, delicate, sharp-tipped, and tapered to a very fine point with a gentle curve. Due to their sharpness and precision, Supercut® Gradle scissors are ideal for removing thin stages during Mohs micrographic surgery and for removing skin tags. Gradle scissors must be used with care, and they should never be used to cut sutures. With improper use, they are quickly dulled, the cutting surfaces are nicked, and the tips are easily malpositioned.

Tissue scissors have relatively short handles and sharp tips. They are available in straight or curved models, with or without the serrations that prevent tissue motion during cutting. Of the various models available, the authors prefer curved Supercut® iris scissors to cut tissue and for sharp dissection. Of the standard tissue scissors available, Supercut® iris scissors have the sharpest edge and they are easily recognized by their black handles. These scissors have a fine bevel angle at the cutting edge, and they are available with smooth edges or one serrated edge. The ‘razor-like’ edge of these scissors enables the surgeon to cut tissue in a smooth, easy motion.

Westcott and Castroviejo scissors are delicate, spring-loaded tissue scissors with very sharp tips (seeFig. 144.9). The configuration of their handle and spring-loaded action make them ideal for manipulation in small delicate sites. For this reason, they are popular with oculoplastic surgeons. They should only be used for cutting thin tissue, such as that encountered in eyelid surgery, or they will dull quickly.

Large, less expensive scissors are sufficient for cutting sutures. Tissue scissors should never be used to cut sutures. Specially designed suture-removal scissors with a half-moon hook on the lower blade are available and the small hooked tip easily grasps the loop and prevents accidental sticks.

Undermining scissors are usually blunt-tipped (for safety) and have longer handles (for comfort). They are available in different sizes to accommodate the various anatomic regions in which skin surgery is performed. Baby Metzenbaum scissors have a high handle-to-blade length ratio, and, with the resultant small blade arc, they have become the most widely used scissor for sharp or blunt undermining (seeFig. 144.12). Larger Metzenbaum scissors are appropriate for extensive undermining in fascial planes on the scalp, trunk and extremities (seeFig. 144.8). Stevens tenotomy scissors and Supercut® Shea undermining scissors are utilized for more superficial, delicate undermining (seeFig. 144.7). The tips of these scissors allow for sharp, less traumatic undermining.

6.1.3 Scissors

Surgeons use surgical scissors during an operation in order to cut tissues at the surface or inside the human body. The blades can be either curved or straight.

The effect of tissue dissection is achieved when the sharpened edges slide against each other when the bows opposite to the joint are closed. For a better wound healing, scissors should cut exactly at the point where the blades meet. Shearing effects due to bluntness or floppy joints have to be avoided. Scissors are usually designed for right-handed persons. High-quality surgical scissors with good tension can also be used by left-handed individuals.

Scissors are the most important and valuable items of the surgical instrument set [2]. They are produced of high grade medical stainless steel, frequently hardened (tungsten carbide). Some approaches have already been made to offer disposable scissors to the market. Up to now, the success is limited [3]. Still today high-quality surgical scissors are reusable. About 2000 different types of surgical scissors are in use (Fig. 6.6).

In visceral surgery, however, the need for highly specialized scissors is low. The most frequently used one is the Metzenbaum type (Fig. 6.7).

For the handling of abdominal tissue and organs which are mostly delicate and fragile, the Metzenbaum scissors are ideally suited. The blades are usually blunt and curved. Since the shanks are comparatively long as compared to the blade, the haptic feedback is excellent. The surgeon feels well whether the structure dissected is soft or hard which helps him to discriminate the border between critical areas (Fig. 6.8A). Even more important is this “sensitivity” of the scissors in case of blunt dissection, since the scissors can also be used to spread the tissue (Fig. 6.8B).

High-quality Metzenbaum shears are expensive and should not be used for too rough tasks, e.g., cutting of material like sutures, meshes, or sponges. For these purposes Mayo scissors are very adequate (Fig. 6.9).

Mayo scissors have semiblunt tips. The blades and handlings are stronger than in Metzenbaum scissors. Often they are called suture scissors or material scissors. Meshes (Fig. 6.10A), plastics, and rubber (Fig. 6.10B) require higher cutting forces than most tissues. The sharp branch of the Mayo scissors facilitates a precise elaboration of the object.

An interesting, relatively new development is so-called “electrical” shears: to prevent smaller bleeding out of capillaries and tiny arteries, specially insulated scissors are available (Fig. 6.11A). Prior to the definitive cut, coagulation of the tissue between the blades is induced by an electrical impulse.

This is usually initiated by the surgeon via a foot pedal. Necessarily, this type of scissors needs two cables which is found irritating by many surgeons (Fig. 6.11B).

Electrical scissors avoid bleeding and speed up the surgical manipulation. However, lateral heat spread has to be considered, which may lead to injuries of adjacent tissue (e.g., nerves). The risk may even be higher than with ultrasound activated dissectors (see Section 6.3: Ultrasound Dissection) [4].

Complications

This is a difficult procedure that is rarely performed; hence, complications from balloon tamponade can be severe and occur in up to 20% of patients.21,25 Major complications include airway obstruction, esophageal rupture, and aspiration pneumonitis. Airway obstruction can be catastrophic and usually results from migration of a dislodged esophageal balloon into the oropharynx.11,26 Prevent proximal migration of the tube by maintaining adequate inflation of the gastric balloon, radiographic confirmation, and periodic monitoring of inflation pressure. In nonintubated patients with a balloon tamponade device, treat respiratory distress as airway obstruction until proved otherwise. In these patients, use surgical scissors to cut across the lumen of the tube just distal to the inflation and aspiration ports. This will result in deflation of both balloons and allow immediate extraction of the device. Given the risk for airway obstruction, always keep surgical scissors at the bedside of patients who have a balloon tamponade device in place.

Esophageal perforation is another catastrophic complication of balloon tamponade that is almost universally fatal. This dreaded complication can occur from a misplaced gastric balloon, an overinflated esophageal balloon, or prolonged inflation of the esophageal balloon and can result in decreased mucosal blood flow, ischemia, and necrosis. To minimize the risk for esophageal perforation, obtain radiographic confirmation of gastric balloon placement before full inflation. In addition, keep the esophageal balloon at the minimum pressure necessary to control hemorrhage. If the device is required for longer than 24 hours, periodically deflate the esophageal balloon to limit mucosal damage and decrease the risk for necrosis.

Aspiration pneumonitis can result from the aspiration of blood, oral secretions, and gastric contents and is a frequent complication of balloon tamponade.27 The incidence of pneumonitis can be decreased by evacuating the stomach and intubating the patient before placement of the tamponade device.22

Additional complications of balloon tamponade include pain; ulceration of the lips, mouth, tongue, and nares; and esophageal and gastric mucosal erosions.27 As discussed, patients with a tamponade device should receive adequate analgesia and sedation. Frequent monitoring of tube placement and pressure can decrease the incidence of esophageal or gastric mucosal erosions.

2.3.2 Surgery for Heart Isolation and Perfusion in the Langendorff Mode

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Prepare a clean surface with surgical scissors, cannula for the aorta (a 18-gauge steel cannula for which the needle is cut diagonally and buffed to remove the sharp edge, 1 in Fig. 2) and the left atrium (a 16-gauge bended cannula; 2 in Fig. 2); a 10 to 15-mm polyethylene catheter (PE-50) with a fluted end (3 in Fig. 2); the pump with the 50-ml syringe containing the cold cardioplegic solution.

Mouse (body weight > 20 g) is anesthetized with sodium thiopental (100 mg/kg ip) or a mixture (1μl/g ip) or a mixture of ketamine (100 mg/ml) and xylazine (20 mg/ml)), heparinized (5000 U/kg sc; to avoid thrombus formation in the aorta and coronary vessels) 15 min prior to the surgery, and fixed in a supine position.

Promptly cut through the abdominal skin and wall using scissors. Once the abdomen is opened, gently pull the liver aside and make an incision on both sides of the thorax (antero-lateral thoracotomy). Immediately chill the exposed heart by flushing of the cold saline solution to stop contraction; flushing is maintained thereafter for the remaining of the procedure.

Rapidly cut through the diaphragm and set the sternum aside. Clear the heart from the pericardium and using forceps carefully remove the thymus and fat around the aorta.

Using a clip, pull a suture wire (black braided silk nonabsorbable surgical suture, 2-0) underneath the aorta using fine surgical forceps and prepare for ligation. Gently pull the aorta between the innominate and the subclavian arteries and practice a hemisection with a 2.5 mm cutting edge scissors. Gently place the cannula into the aorta and make a ligature on the cannula. Finally, cut and remove heart and lungs as a whole by cutting across the aorta, vena cava, pulmonary veins, and trachea.

Slowly insert into the aorta an 18-gauge cannula (1 in Fig. 2), which is connected using polyethylene tubing (75 cm length) to the 50-ml syringe containing cold (4 °C) cardioplegic solution. (N.B. Be careful not to force the cannula beyond the aortic valve.) When the cannula is correctly positioned into the aorta, the coronary arteries become translucent. The cardioplegic solution is retroperfused into the large vessels and the blood will be flushed. The cannula is fixed onto the aorta using a double knot (Fig. 2).

The heart is then excised by slowly cutting underneath tissues, but keeping part of the lungs to preserve the integrity of the left atrium. While maintaining cannula in the aorta–heart axis, quickly place the cannula with the heart onto the setup, on the three-way valve which is connected to the compliance chamber and the Langendorff line. Prior to positioning of the heart, allow buffer from reservoir 1 to flow through the system to avoid formation of air bubbles, which could clog the coronary arteries. The heart should start beating within a few seconds upon perfusion with buffer at 37 °C.

While the heart is being perfused in the retrograde or Langendorff mode (Fig. 1, green arrow) via the aorta (Fig. 2), carefully and quickly remove remaining lung and fat tissues from the heart, while preserving the integrity of the left atrium, in order to have access to the pulmonary veins.

Equipment

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Sterile gown, gloves, drapes, and any other equipment necessary for the clinician to follow universal blood and body fluid precautions.

Povidone–iodine solution (or chlorhexidine if patient allergic to iodine).

Blunt-tipped straight surgical scissors (e.g., Mayo scissors; a scalpel may also be used but the editors do not recommend this because it can cause significant injury to the mother, fetus, or clinician).

Needle holder.

Nontraumatic forceps.

Vaginal retractor(s).

Ring forceps.

4 × 4 sterile gauze sponges.

2-0 or 3-0 polyglycolic braided absorbable sutures (e.g., Vicryl, Dexon, Polysorb; preferred over chromic, which is associated with an increased risk of episiotomy breakdown and more discomfort during the first 3 days of healing; however, polyglycolic sutures may more frequently cause local irritation and work themselves to the skin surface or fail to heal and have to be removed weeks later) on a large, curved cutting or tapered-point needle. Rapidly absorbable versions of polyglycolic sutures are now also available and may decrease the need for suture removal within the first 3 months after episiotomy repair.

Allis clamps (especially for third- and fourth-degree extension repairs).

4-0 or 5-0 polyglycolic suture on smaller curved needle (for fourth-degree extension repairs).

Note: If effective regional (e.g., epidural, pudendal) anesthesia is not in place, a 10-mL syringe with a 1.5-inch, 27-gauge needle should be available to locally infiltrate the anesthetic of preference (usually 10 mL 1% lidocaine without epinephrine; use ≤30 mL lidocaine).

3.2 Final sacrifice for tumor endpoints

3.2.2 Final sacrifice endpoint

The mice are typically sacrificed between 20 and 30 week. At 20 week., small adenomas exist and are visible on the lung surface, while at 30 week., the tumors are significantly larger and some adenocarcinomas are present. Earlier time points can also be used to assess smaller adenomas, hyperplasias, and the inflammatory microenvironment of the tumor bearing lungs. The manner in which the tumors are analyzed depends on the equipment available. If a micro-CT is available, then assessing tumor development from weeks 8–30, would be a valuable tool.

A.

For the final sacrifice of tumor analysis, mice are weighed and euthanized with 120 mg/kg Fatal Plus solution (MWI) or whatever accepted method of euthanasia is preferred (see Note 4.6).

B.

At this point, the lung tumors can be counted for in fixed tissue (see below) or unfixed tissue (Section 3.2.4). There is an alternative to these two choices (see Note 4.7).

1.

For lung fixation, the mice can be fixed in several ways for surface tumor analysis. (1) Using Tellyesniczky's fixative that turns the tumors bright white and the lung opaque (see Note 4.8) or (2) in 10% NBF. Unless the lungs are processed for other endpoints, it is best to use the entire lung for counting tumors. If one of the goals is to do a lot of immunostaining, the best fixative is NBF; Tellyesniczky's fixative is not good for epitope retrieval for most targets. Tellyesniczky's fixative likely turns the tumors bright white due to the acetic acid in the fixative. Acetic acid causes the color change due to the increased percentage of abnormal nuclear protein and increased number of dysplastic cells and is used in colposcopies for cervical cancer detection (https://screening.iarc.fr/colpochap.php?lang=1&chap=4).

2.

Prior to fixation, carefully perfuse the lung through the heart with sterile saline using a 5 mL syringe to remove the blood followed by careful and slow inflation of the lung with fixative via trachea (~ 1 mL/25 g body weight). To do so, make a 1.5–2 mm small longitudinal incision in the trachea using sharp scissors on the ventral side of the neck. Insert a 20–24 g plastic catheter (or metal stubber needle) into the trachea about 0.5 cm. Ensure that the catheter is not inserted too far down into the trachea, as this can lead to damage of the lung structure. After the lung is inflated based on body weight (~ 1 mL/25 g body weight), tie off the trachea.

3.

The lung is then carefully removed en bloc and submersed into fixative for 24 (NBF) or 48 h (Tellyesniczchy's). For the NBS, the lung is transferred to 70% ethanol after 24 h of fixation (to maintain epitope integrity) and for Tellyesniczchy's, the same after 48 h.

4.

After at least 48 h, each lung lobe is carefully dissected apart from the other using a fine-tipped forceps and kept in the fixative prior to counting (see Fig. 2).

Fig. 2. Lung lobes of mouse dissected apart to count tumors. Lungs were fixed in Tellyesniczky's fixative. Red arrows indicate tumors on the surface of the lung. These are most likely adenomas.

C.

To evaluate the surface tumors between 20 and 30 week., a dissecting microscope is used with careful analysis. Each lobe is carefully counted for tumors and each tumor is sized at the same time using a digital caliper. Tumors less than 200 μm are typically considered microadenomas if they are round and visible. However, this is subjective. Hyperplasias and dysplasias follow the normal lung architecture.

1.

All tumor counts from the five lobes are combined and considered tumor multiplicity for that mouse.

2.

Then, all tumor numbers per mouse are averaged to determine the tumor multiplicity differences between treatments (i.e., vehicle control, MCA, BHT, and MCA/BHT).

3.

Tumor sizes are determined using a digital caliper. For spherical lesions, the calculations for tumor volumes use (4/3 π)(r3) and for non-spherical lesions (Length × Width × Height).

4.

The lungs are then processed and sectioned for histology, typically at a histology core.

2.2.1 Embryo culture

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Two pairs of watchmaker's forceps #5 (Roboz, RS-4978) and small surgical scissors (Roboz, RS-5910) for embryo dissection

3.5, 6, and 10 cm plastic Petri dishes (Falcon, 351001; 351007 and 351029)

Organ culture dishes (Falcon, 353037)

3.5 cm glass-bottom dishes (MatTek, P35G-1.5-14-C) or LabTek coverslip bottom chambers (NUNC, 155360; 155379; 155382)

CoverWell perfusion chamber gaskets, 9 mm diameter; 1.0 mm deep (Invitrogen/Molecular Probes, C18140) or 2.0 mm deep (Invitrogen/Molecular Probes, C18141)

Mouth pipette (homemade) consistent of a mouthpiece (HPI Hospital Products Med. Tech., 1501P-B4036-2), latex tubing (latex 1/8 in. ID, 1/32 in. wall, Fisherbrand, 22362772), and very fine pulled glass Pasteur pipettes using a 1000 μl pipette tip connector

Plastic transfer pipettes (Fisherbrand, 13-711-7M) to transfer embryos stages E7.5 and older

Pulled Pasteur pipettes

Suction holding pipette (optional; Eppendorf CellTram Air, 5176000.017)

1 ml syringe and 26-gauge needle (Becton Dickinson, 309623), 30-gauge needle with a blunt end (Becton Dickinson, 305106). Use sandpaper or sharpening stone for blunting the needle.

Embryo tested light weight mineral oil (Sigma, M8410)

Incubator providing a humid atmosphere and constant level of 5% CO2

Roller apparatus (BTC Engineering, Cambridge, UK)

Industrial gas supply containing gas mixtures of 5% CO2/95% O2; 5% CO2/20% O2, or 5% CO2/5% O2

Microscope with an environmental chamber to keep temperature and gas levels stable throughout the culture

Human eyelashes, or cat whiskers, sterilized with 70% ethanol. (Note: To prevent animal cruelty, cats should not be harmed during whisker collection. We therefore recommend the use of cat whiskers which have been naturally shed by the animal.)

Subcutaneous Implantation

1.

The animal should be anesthetized and prepared for surgery following institutional guidelines.

Using sterile surgical scissors, make a 5- to 10-mm vertical skin incision on the flank.

Insert a straight forceps gently 2 cm into the SC space to create a pocket large enough for a tumor fragment.

Using sterile straight forceps insert the tumor fragment previously prepared.

Close the skin incision using sutures, clips, or tissue glue.

Animals should be provided postoperative care and monitoring according to institutional guidelines.

2.1 Ad hoc probe introduction: Injection, electroporation

Due to their accessibility, muscles of the mouse distal extremities are particularly amenable to probe injection and electroporation (Figs. 19.1A and B and 19.2A–G). On the one hand, injected probes might be cell-permeant or cell-impermeant, such as tetramethyl rhodamine methyl ester (TMRM) (a mitochondrial membrane potential marker; Romanello et al., 2010) or α-bungarotoxin coupled to chromophores (marking AChRs in neuromuscular junctions; Röder et al., 2008, 2010), respectively. On the other hand, electroporation (Figs. 19.1B and 19.2A–G) is a quick and efficient method to introduce cDNAs for expressing genetically encoded fluorescent probes or other genes or siRNAs, which modulate any tissue function of interest. As shown previously, transfected cDNAs remain actively transcribed and translated for up to 4 weeks after electroporation in skeletal muscle (Dona et al., 2003). This is most likely due to the postmitotic status of adult skeletal muscle. Although very high levels of up to 80% of transfected fibers might be reached for some constructs, expression of heterologous proteins being targeted to specific sites or organelles, such as the synaptic membrane, mitochondria, or the sarcoplasmic reticulum can be much more cumbersome. This is particularly true for transmembrane proteins of the synapse, since the secretory pathway mediating the delivery of synaptic proteins is scarcely developed in skeletal muscle. For two main reasons, we usually wait a couple of days (≥ 10) before imaging transfected muscles. First, a mild, unavoidable inflammation typically occurring at the site of electroporation is completely healed by then. Second, many proteins in skeletal muscle exhibit low turnover rates, such as the AChR, which has a half-life of > 10 days. It is also critical to have always the same, experienced persons performing the electroporation, because this strongly reduces damaging of muscle, which is almost zero in the optimal case, but can be problematic upon inadequate performance. It is worth noting that electroporation efficiency might vary from mouse strain to mouse strain and it is particularly low in muscles with a lot of fat and connective tissue, such as in dystrophic animals.

Of the different protocols available for muscle transfection, we prefer to use the method from Dona et al. (2003), where the transfected muscle (e.g., tibialis anterior (TA)) is exposed by simple surgery (Figs. 19.1A, B and 19.2A–G). This is ideal for local and muscle-specific transfection and allows transfecting either the whole muscle (by injecting at different sites) or only superficial fiber layers (sufficient for imaging). For transfecting lower hindlimb muscles, such as TA or extensor digitorum longus (EDL), we use spatula electrodes of 1 mm thickness and 4 mm width (Fig. 19.2D). These are connected to a BTX ECM830 square pulse generator (Fig. 19.2A). After orienting the anesthetized mouse on the side to be transfected, the lower hindlimb is stretched over a pillow and crepe tape is used to fix the foot (Fig. 19.2B). After shaving the leg, a longitudinal cut of ∼ 6 mm length opens the midline of the lower hindlimb exposing a view on the whitish border between anterior and posterior muscles (Fig. 19.2C). The shiny fascia (see Fig. 19.2J) enclosing this whole arrangement is then opened by a second longitudinal cut. This allows separating anterior from posterior muscles and seeing the tibia. Now, a closed pair of small surgical scissors penetrates between the tendon of the TA and the tibia. Careful opening of the scissors creates a hole through which the electrode can be inserted (Fig. 19.2D). After fixing the electrode on the pillow with tape, a Hamilton syringe is used to slowly inject up to 20 μl of cDNA in physiological solution (less is better, Fig. 19.2E). Depending on the construct, 0.5–20 μg of cDNA might be used. Make sure that no liquid is leaving the injection site, because this usually results in low transfection efficiency. If necessary, additional, cell-impermeant labels should be injected subsequently. Then, carefully place the second electrode on top of the muscle (Fig. 19.2F) and apply pulses in the following pattern: five pulses with 5 ms duration each are given at an interval of 200 ms between pulses. Voltage is typically 50 V/cm of muscle. Make sure to adapt the voltage to muscle thickness. Then, carefully remove both electrodes and close the skin with two to four surgical stitches (Fig. 19.2G), followed by sterilizing the wound. Experienced operators perform a transfection in a few minutes, and animals immediately walk, jump, and behave normally after complete recovery from anesthesia. To help fast recovery, let the animals wake up on a heating plate at 37 °C. Animals, which do not behave normally few hours after transfection, suffer from pain or infection and should additionally be treated with an antibiotic and analgesic.

Instruments and Materials Used for Generation of Slice Explant Cultures

The following instruments are routinely used during the dissection and plating procedures: dissecting microscope, fiber optic lighting, tissue chopper, repeating Eppendorf pipettes, small oven, large surgical scissors (decapitation), small scissors (to remove the brain from the calvarium), fine-tipped forceps (removal of the pia, blood vessels, etc., and manipulation of the tissue slices), polished flat-surfaced small spatulas (transfer of the blocked tissue and tissue slices), razor blade holders and breakable blades (to dissect out the area of interest), and aclar film (plastic disks used on tissue chopper, which can be sterilized and discarded after using).