Clinical Training 2 End of Practice written exam content-1.pdf

JORDAN UNIVERSITY OF SCIENCE AND TECHNOLOGY Faculty of Applied Medical Sciences Department of Allied Health Sciences Respiratory Therapy Program Clinical Training 2 - (RTH386) End of Practice Written Exam Content 1  Arteri...

JORDAN UNIVERSITY OF SCIENCE AND TECHNOLOGY Faculty of Applied Medical Sciences Department of Allied Health Sciences Respiratory Therapy Program Clinical Training 2 - (RTH386) End of Practice Written Exam Content 1  Arterial blood gas (ABG) Sampling Arterial blood gas (ABG) analysis is a beneficial diagnostic test for assessing ventilation, acid-base status, and oxygenation. Collection of an arterial sample may be done quickly, and it provides important information for decision-making in the management of the patient requiring oxygen or ventilatory assistance. Indications for blood gas and pH analysis and hemoximetry 1. the need to evaluate the adequacy of a patient’s ventilatory (PaCO2), acid-base (pH and PaCO2), and oxygenation (PaO2 and O2Hb) status & the oxygen-carrying capacity. 2. the need to quantify the response to therapeutic intervention (e.g., supplemental oxygen administration, mechanical ventilation) and/or diagnostic evaluation. 3. the need to monitor the severity and progression of documented disease processes. Contraindications to performing pH-blood gas analysis and hemoximetry 1. an improperly functioning analyzer. 2. an analyzer that has not had functional status validated by analysis of commercially prepared quality control products or has not been validated through participation in a proficiency testing program(s). 3. a specimen that has not been properly anticoagulated. 4. a specimen containing visible air bubbles. 5. a specimen stored in a plastic syringe at room temperature for longer than 30 minutes, stored at room temperature for longer than 5 minutes, or stored at room temperature in the presence of elevated white blood cells or platelet count (PaO2 in samples drawn from subjects with very high leukocyte counts can decrease rapidly. Immediate chilling and analysis is necessary). Hazards/Complications Possible hazards or complications include: 1. infection of specimen handler from blood carrying the human immunodeficiency virus, or HIV, hepatitis B, and other blood-borne pathogens. 2. inappropriate patient medical treatment based on improperly analyzed blood specimen or from analysis of an unacceptable specimen or from incorrect reporting of results. Monitoring Monitoring of personnel, sample handling, and analyzer performance to assure proper handling, analysis, and reporting should be ongoing, during the process. CBGs ( Capillary Blood Gases) Assessment of Need Capillary blood gas sampling is an intermittent procedure and should be performed when a documented need exists. Routine or standing orders for capillary puncture are not recommended. The following elements may assist the clinician in assessing the need for capillary blood gas sampling: 1. History and physical assessment. 2. Noninvasive respiratory monitoring values as : 2 A. Pulse oximetry. B. Transcutaneous values C. End-tidal CO2 values 3. Patient response to initiation, administration, or change in therapeutic modalities. 4. change in therapeutic modalities. 5. Lack of arterial access for blood gas sampling. CBGs Monitoring 1. FIO2 or prescribed oxygen flow. 2. Oxygen administration device or ventilator settings. 3. Free flow of blood without the necessity for “milking” the foot or finger to obtain a sample 4. Presence/absence of air or clot in the sample. 5. Patient temperature, respiratory rate, position or level of activity, and clinical appearance. 6. Ease or difficulty of obtaining sample. 7. Appearance of puncture site. 8. Complications or adverse reactions to the procedure 9. Date, time, and sampling site. 10. Noninvasive monitoring values: transcutaneous O2 & CO2, end-tidal CO2, and/or pulse Oximetry. 11. Results of the blood gas analysis. Anatomical Locations For Arterial Puncture The radial artery’s location makes it easily accessible. It is located in the wrist on the radial side (thumb side), close to the surface of the skin. It is the site most commonly used for taking a patient’s pulse. A big advantage of performing arterial puncture at the radial site is the safety afforded by the presence of collateral circulation. The hand is supplied with blood by both the radial and ulnar arteries. If circulation is inadvertently interrupted resulting from radial artery puncture, the ulnar artery will continue to supply the circulatory needs of the hand. There are no veins or nerves immediately adjacent to the radial artery; consequently, arterial sampling at this site is facilitated by a reduced chance of inadvertent venous puncture or nerve damage. The disadvantage of radial artery puncture is the small size of this artery. The radial artery is a small target. However, through careful observation, palpation, and considerable practice, the radial artery can be punctured easily. However, in case of hypotensive and hypovolemic states or low cardiac output, puncture at this site may be particularly difficult. Brachial Artery The site where the brachial artery is commonly punctured is at the elbow in the antecubital fossa. It is located on the medial side of the fossa near the insertion of the biceps muscle at the radial tuberosity. An advantage of the brachial artery puncture site is its size. It is large and easily palpated. There are several disadvantages to using the brachial artery. It is close to both a large vein and a nerve. Inadvertent venous sampling is common at this site. Accidental contact with the nerves at this site may cause extreme discomfort. Also, this site does not have the advantage of collateral circulation. Inadvertent injury leading to the stoppage of circulation may result in the loss of the limb. Femoral Artery The femoral artery is accessible for arterial sampling in the groin. It may be palpated laterally from the pubis bone. The femoral artery is very large. It is easily palpated and presents a large target. The femoral artery may be the only site where arterial sampling is possible in cases of hypovolemia or hypotension, during cardiopulmonary resuscitation (CPR), or with low cardiac output. There are several disadvantages 3 to arterial puncture at this site, including the proximity of a major vein and a lack of collateral circulation. The artery may also be deep and difficult to locate. Atherosclerotic plaques commonly form in the femoral artery. If a plaque is dislodged as a result of arterial puncture, circulation to the entire leg may be compromised by the formation of emboli. In addition, the proximity of the femoral vein makes the certainty of arterial sampling questionable. The Modified Allen’s Test for Collateral Circulation An advantage of performing an arterial puncture at the radial site is that the vascular anatomy allows testing for collateral circulation. The modified Allen’s test should be done before arterial puncture to determine the adequacy of circulation supplied by the ulnar artery. To perform the modified Allen’s test, elevate the patient’s hand higher than the level of the heart. Have the patient make a fist for approximately 30 seconds. Apply pressure to both the radial and ulnar arteries occluding them. Have the patient open his or her hand (it should appear blanched). Release pressure on the ulnar artery. Color should return to the hand within 7 to 10 seconds. If the hand does not flush pink, it is likely that blood flow through the ulnar artery is insufficient to provide circulation if the radial artery loses patency. If this happens, try the other hand. If this also fails, choose the brachial site for puncture. This test may be done on an unconscious patient by holding the hand above the heart for 30 to 60 seconds before releasing pressure on the ulnar artery. 4 The modified Allen’s test: (1) The radial and ulnar arteries are palpated. (2) The patient clenches the fist, and both arteries are compressed manually. (3) The patient opens the fist (note the blanching of the palm). (4) The ulnar artery pressure is released; if the Colour returns to pink, the test is positive; if the Colour does not return to pink, the test is negative. BLOOD GAS SAMPLING ERRORS The reliability of blood gas analysis is very technique-dependent. Every step of the process, from the preparation of equipment to reporting the data, has potential problems that affect data reliability. That is, knowledge and skill as a respiratory care practitioner will often determine the accuracy of the procedure. Knowledge of the factors that contribute to sampling errors will help to prevent their occurrence in clinical practice. If a sample is questionable, the relevant facts should be noted with the results of the analysis. Clinical decisions are frequently based on data assumed to be entirely accurate. Factors that contribute to sampling errors Bubbles If the sample is aspirated, air bubbles are often present in the collected blood sample. These bubbles must be expelled immediately upon collection. Room air contains enough oxygen that it can diffuse into the sample, increasing the arterial oxygen tension (PaO2), or if the PaO2 is greater than 160 mm Hg, it can decrease the PaO2. This diffusion problem is especially true in patients with hypoxemia. Carbon dioxide is present in the atmosphere in a concentration of only 0.003%, or around 2 mm Hg at sea level. Because arterial blood normally has a carbon dioxide tension (PaCO2) ranging from 35 to 45 mm Hg, CO2 dissolved in blood will tend to diffuse into the bubbles in the sample, lowering the measured value. Ideally, if a large quantity of bubbles is present in the sample, it is best to discard it and draw another sample, being more attentive to the technique. However, this may not be practical in the clinical setting. results can change dramatically (American Association for Respiratory Care, 2019). Immediately after collection, cool the sample in a slush of ice and water. Ice slows the cells’ metabolism, and even a delay of 30 minutes will not significantly affect the analysis results. It is best, however, to analyze the sample as 5 soon after collection as possible. Ten minutes is optimal. Samples held longer than this may show lower PaO2, higher PaCO2, or a pH less than the patient’s actual pH. Use of the Proper Anticoagulant Oxalates, ethylene diamine tetra-acetic acid (EDTA), and the citrate anticoagulants available will alter the pH of the arterial sample (American Association for Respiratory Care, 2020). Sodium heparin is the best anticoagulant to use in arterial blood sampling. You need 0.05 ml of sodium heparin to anticoagulated 1 ml of blood, if too much is used (more than 0.1 mL of heparin per 1 mL of whole blood), will cause acidosis in the blood sample. If preparing a glass syringe, a safe general rule to follow is to expel all excess heparin. Heparin will be left in the needle and needle hub, occupying a minimal volume. The syringes in blood gas sampling kits often contain crystalline heparin or lithium heparin. No aspiration of additional anticoagulant is necessary. Simply draw the sample, and the anticoagulant will dissolve. However, it is important to mix the sample by gently rolling or shaking the syringe after collection. Venous Sampling In a sample drawn from a patient with hypoxemia, it is difficult to distinguish arterial blood from venous blood by color. When an arterial sample is drawn, the plunger of the syringe tends to pulsate as the sample fills the barrel. If the syringe does not fill without assistance, be suspicious of the sample site. Patients in cardiopulmonary arrest, hypovolemia, hypotension, or low cardiac output often have low blood pressure. Samples from these patients must usually be aspirated. Drawing from the brachial or femoral site will help to ensure obtaining an arterial sample. In the event of collection of a venous sample, draw another sample for analysis. If this is not possible, note with the sample results that it may be a venous sample. Patient Anxiety It is a rare person who enjoys having blood drawn. It is natural to be a little apprehensive before the skin is punctured by a needle. However, if extreme, this anxiety may lead to hyperventilation and consequent altering of the PaCO2. Anxiety can be minimized by doing the procedure quickly and being prepared before reaching the bedside. Do not stress the pain and discomfort a patient may experience with arterial puncture. CAPILLARY BLOOD GAS SAMPLING In infants, capillary blood gas sampling is frequently performed instead of arterial puncture. Performing arterial puncture in an infant requires a large degree of skill and good technique for best results. The infant’s vessels are very small and are difficult to palpate and puncture. Technique Capillary samples are usually obtained from the infant’s heel but may also be obtained from the finger. When capillary blood is drawn from the heel, this sampling technique is sometimes referred to as a heel stick. Arterialization (warming to maximize blood flow) of the infant’s heel is done before sampling, and then a lancet is used to puncture the skin surface. A sample from an adequately arterialized limb will yield reliable pH and PaCO2 values, whereas PaO2 values will vary from those determined using blood drawn by the arterial puncture. 6 Capillary Sampling Errors Poor Blood Flow In performing capillary blood sampling, it is important to obtain a free-flowing sample. Do not squeeze the infant’s foot when drawing the sample into a capillary tube. Squeezing the foot excessively could result in injury to the foot, leading to altered blood gas values. If blood at the puncture site does not flow freely, repeat the puncture to obtain a freely flowing blood sample. Introduction of Air into the Sample As with arterial puncture techniques, it is important to minimize the blood sample’s exposure to air. Air may be introduced when it is drawn into the capillary tube, causing bubbles. The presence of air bubbles will alter the blood gas results; therefore, the introduction of air should be avoided. If the sample contains visible bubbles, discard it and draw another (or remove the air bubbles as they enter the capillary tube). Inadequate Mixing of Heparin Once the sample has been drawn into the capillary tube, one end may be sealed with clay or a rubber stopper. A small metal rod is inserted into the capillary tube, and a magnet is passed back and forth along the length of the tube to mix the heparin in the capillary tube with the blood sample. It is important to mix the heparin well to avoid clotting before sampling. SUPPLIES NEEDED FOR ARTERIAL PUNCTURE The supplies needed for arterial puncture are usually contained in an ABG kit. If the practitioner needs to assemble supplies separately, the supplies needed are listed in this table 1 5-mL pre-heparinized disposable sampling 6 A plastic bag or another container to syringe transport the sample 2 Needles (20 to 25 gauge, in various lengths) 7 Ice slush 3 Rubber stopper or rubber syringe cap 8 Lidocaine anesthetic 2% solution (if ordered) 4 Adhesive strip or Elastoplast tape 9 Disposable latex gloves 5 Iodine and alcohol prep pads 10 Eye protection (goggles or face shield) 7 Patient-Related Considerations A physician’s order is required before performing this or any other procedure. It is important to check the patient’s chart for a physician’s order for anticoagulant therapy or oxygen therapy before arterial sampling. Anticoagulant therapy may necessitate putting pressure on the site for up to 15 minutes to stop the bleeding following an arterial puncture. It is important to assess if the patient is receiving the proper oxygen therapy (or to identify the lack of therapy) before doing the puncture. ABG analysis is a useful tool for judging the adequacy of oxygen therapy. If blood is drawn while the patient is receiving the wrong oxygen regimen, however, a repeat arterial puncture is necessary. Mistakes can be avoided by checking the chart and the patient first. Usually, 10 to 30 minutes are needed before arterial puncture after any oxygen concentration change. Use of an Anesthetic The use of an anesthetic before arterial puncture requires a physician’s order and a specific protocol. The puncture site should be prepared as though an arterial puncture is being performed. Draw up 0.1 to 0.2 mL of 2% lidocaine into a tuberculin syringe with a 25-gauge needle. Puncture the skin near the artery and draw back on the plunger. If blood appears in the syringe, the needle is in a vessel. Withdraw the syringe and redirect the needle. Repeat the aspiration step. If there is no blood return, make a small wheal by injecting part of the anesthetic. Try to surround the artery with an anesthetic. Allow 2 to 3 minutes to lapse before performing the arterial puncture. Puncture Preparation Prepare the syringe as outlined in the previous section. If using a pre-heparinized syringe, follow the manufacturer’s instructions for preparation. It is best to arrange all of the supplies needed within easy reach. The rubber stopper, 2 *2-inch gauze pads, container with ice slush, and adhesive strip should be arranged so that immediate retrieval is possible. If doing a radial puncture, perform the modified Allen’s test. Carefully palpate the puncture site. Try to form a mental image of the course and direction of the artery. Note the strength of the pulse and try to estimate the depth of the artery below the skin. Using an iodine-based prep pad, cleanse the site. Use firm pressure, scribing a circle from the puncture site out. Follow the iodine-based preparation with an alcohol prep pad, using the same technique. Obtaining the Specimen Radial and Brachial Sites: Hold the syringe like a pencil. Palpate the pulse and visualize the artery location. For sampling at the radial site, with the needle bevel up, puncture the skin at a 30°-45° angle. Once the needle is below the skin surface, visualize the artery location and slowly advance the needle toward the artery. Observe the hub of the needle. When the artery is punctured, blood will quickly appear in the needle hub. This is termed a flash. Upon seeing the flash, do not move. Allow the syringe to fill. For sampling at the brachial site, with the needle bevel up, puncture the skin at a 45° to 90° angle. Advance the needle slowly toward the artery, watching for the flash. When the flash is observed, do not move. Allow the syringe to fill. Femoral Site: The femoral artery is deep and may require a longer needle for successful puncture. With the bevel of the needle facing the patient’s head and perpendicular to the skin’s surface, puncture the skin. Watch for the flash and allow the syringe to fill. 8 Post-puncture Care After the sample is collected, withdraw the needle and apply firm pressure with a 2 *2-inch gauze pad. Expel any air and insert the needle into a rubber stopper to seal it. Many disposable blood gas collection kits come with one-handed safety caps. The use of these devices minimizes the risk of inadvertent needle sticks by elimination of recapping the syringe. Continue to apply firm pressure to the puncture site for a minimum of 5 minutes. While holding firm pressure to the puncture site, mix the sample in the syringe. The sample may be mixed by gently rolling the syringe between the thumb and forefinger. Rolling the syringe will mix the heparin with the blood and prevent clotting. Ice the sample after mixing by placing it into the container of ice slush. Check the puncture site after 5 minutes has passed. Observe the color of the skin distal to the puncture. Check for circulation by palpating the artery distal to the puncture site. The skin should be warm to the touch, and when the tissue is firmly pressed and released, capillary refill should be evident. An adhesive strip may be applied now. Label the sample with the patient’s name and room number, time of collection, and the fraction of inspired oxygen (FIO2) inhaled by the patient, respiratory rate, or ventilator settings as appropriate. Transport the sample to the laboratory for analysis. After 20 minutes, check the puncture site again, using the same criteria described earlier. Table 1: Necessary Components of Arterial Blood Gas Results Component Definition Normal values H+ Hydrogen ions, inversely proportional to pH 35-45 mmol/L pH Acidity/alkalinity 7.35-7.45 PaO2 Partial pressure of oxygen in arterial blood 80-100 mmHg SaO2 Arterial oxygen saturation 95-100% PaCO2 Partial pressure of CO2 in arterial blood 35-45 mm Hg HCO3- Bicarbonate in blood 22-26 mEq/L BE Base excess (amount of excess or –2 to +2 mmol/L insufficient amount of base in blood) –ve in acidosis, +ve in alkalosis. 9  Table 2: Arterial versus venous blood gas Component Arterial blood average/range Mixed venous average / range pH pH 7.40 (7.35-7.45) 7.36 (7.31-7.41) PaO2 80-100 mmHg 35-40 mmHg O2 saturation 94-95% 70-75% PaCO2 35-45 mmHg 41-51 mmHg HCO3- 22-26 mEq/L 22-26 mEq/L BE –2 to +2 –2 to +2 Table 3: Grading of hypoxemia Severity SaO2 PaO2 Mild 90-94mmHg 60-79 mmHg Moderate 75-89 mmHg 40-59 mmHg Severe 10% chance of difficult airway O – Obstruction – Is there a tumor, epiglottitis, or recent neck surgery? N – Neck mobility – Is the patient in a cervical collar, are they elderly? Mallampati Score: 23 Equipment and preparation: 1. Identify the component of the endotracheal tube: Equipment selection and testing: Gather the necessary equipment: a. Batteries. b. Laryngoscope handles and blades c. Bulbs d. Endotracheal tube e. 10-ml syringe f. Cuff inflator 2. Assemble the laryngoscope (install batteries). 3. Select the appropriate blade type and size (3 and 4 are considered adult sizes) 24  Types of blades : Figure 1 Macintosh blade Figure 2 Miller blade. 4. Attach the blade and observe whether the light is white, tight, and bright. 5. Select the appropriate tube type and size. Sizes range from 2.5 to 9.0 mm in inside diameter and 12 to 32 cm in length. In general, a woman is intubated with a 7 or 7.5 orotracheal tube, and a man is intubated with an 8.0 or 8.5 orotracheal tube. Always have three sizes ready, half size smaller and half size larger. 6. Using the cuff inflator inject the air into the pilot balloon and observe the cuff inflation and pressure. 7. Ensure that pressure does not decline, deflate the cuff completely Endotracheal tube Insertion 1. Wash your hands and apply standard precautions: use of gloves, gown, and face shield or mask and goggles is necessary 2. Open the endotracheal tube package halfway, leaving the cuffed end aseptically protected in the wrapper. Insert a stylet into the tube, 3. Make sure that the tip does not protrude from the other end. Shape the tube so that the curve is maintained. 25 4. Using the manual resuscitator (bag-valve-mask), pre-oxygenate the patient with 100% oxygen. 5. Remove the oropharyngeal airway if one is present. 6. Position the head in the sniffing position by flexing the neck and tilting the head backward. Do not hyperextend. 7. Open the mouth using the crossed-finger, or scissor technique. 8. Holding the laryngoscope in your left hand. Insert the blade into the mouth 9. Pushing aside the tongue to the left advance the blade until the epiglottis is visualized. The epiglottis is displaced indirectly by advancing the tip of the blade into the vallecula. The epiglottis is displaced directly by advancing the tip of the blade over its posterior surface. 26 10. Continue to advance the tip of the blade into the Vallecula, and indirectly expose the glottis by applying an upward and forward lift with your wrist kept straight. Do not use the blade as a lever and rest it on the upper teeth. 11. Sellick’s maneuver, the application of pressure on the cricoid cartilage by an assistant, is sometimes beneficial in visualization 12. After you have seen the ET tube cuff pass roughly 1/2″ beyond the vocal cords: −Gently remove the blade −Secure tube with right hand −Remove the stylet from the tube 13. Inflate the distal cuff with 5 to 10 mL of air, then detach the syringe from the inflation port. No more than 30 seconds should be devoted to any intubation attempt. If intubation fails, immediate ventilation and oxygenation of the patient for 3 to 5 minutes before the next attempt should occur. 14. Ventilate and oxygenate the patient. Observe for bilateral symmetrical chest expansion. 15. Auscultate for bilateral breath sounds. Auscultate the epigastric region to listen for air in the stomach. 27 16. If no breath sounds are heard, deflate the cuff, remove the tube, and ventilate with a bag-valve mask until the procedure is attempted again. 17. If unilateral sounds are heard, deflate the cuff and withdraw the tube gently while continuing to bag the patient until bilateral sounds are heard. Then re-inflate the cuff. 18. Secure the tube with tape or with a commercial ETT holder. Securing ETT with tape commercial ETT holder 19. Attach the patient to an oxygenation or ventilation device. 20. Remove your gloves and other personal protective equipment and wash your hands. Small and Large Volume Nebulizer An aerosol is defined as a suspension of solid or liquid particles in a gas. Aerosols may be administered to deliver medication or to deliver a bland solution for the purpose of sputum induction, treatment of upper airway edema, or humidification of a bypassed or compromised upper airway. Many devices are available to generate aerosols for application to the upper or lower airway, including small-volume nebulizers (SVNs), and large-volume nebulizers (LVNs). These devices are commonly used to administer bronchodilators, corticosteroids, and other medications to treat conditions such as asthma, chronic obstructive pulmonary disease (COPD), and cystic fibrosis. They work by converting liquid medication into a fine mist that can be inhaled directly into the lungs, providing targeted therapy to the affected airways. The selection and use of the most appropriate device is based on the specific clinical application and the desired therapeutic goals. You should consider several factors in this selection, including the intended target area of the respiratory tract to be treated; and patient preference, coordination, tolerance, and cooperation. 28 Large Volume Nebulizer (LVN) LVN is the most common device used to generate bland aerosols. These devices are pneumatically powered, attaching directly to a flowmeter and compressed gas source. A variable air entrainment port allows air mixing to increase flow rates and alter FiO2 levels. A) Identify the component of LVN: 29 B) Identify the aerosol delivery devices: C) Application of an aerosol-generating device (LVN): 1. Verify the physician’s order or protocol for mode of delivery and Fio2. 2. Gather the necessary equipment: Large-volume nebulizer, prefilled if available Corrugated tubing Sterile water Heating element Temperature probe and in-line adaptor 30  Oxygen flowmeter and gas source Water trap or aerosol T-drainage bag Scissors Appropriate aerosol delivery device (mask, T-piece, tracheostomy collar, or face tent) 3. Wash or sanitize your hands. Apply standard PPE. 4. Introduce yourself and verify the patient’s identification. Explain the procedure to the patient. 5. Assemble the equipment. 6. Adjust the gas source to the appropriate flow rate for adequate flow to meet the patient’s inspiratory demand. In most cases, this is between 8 and 12 Lpm. 7. Attach the delivery device to the patient and ensure patient comfort. 8. Analyze the Fio2 and adjust the entrainment selector if necessary. 9. Document the procedure appropriately. Small Volume Nebulizer (SVN) A small-volume nebulizer (SVN) is a medical device used to deliver medication in aerosol form to the lungs for respiratory therapy. It consists of a nebulizer chamber, tubing, and a mouthpiece or mask. Medication is placed into the nebulizer chamber, where it is aerosolized into fine droplets by compressed air or oxygen. A) Identify the component of SVN 31 B) Categories of SVN 32 C) SVN delivery: 33 D) SVN Administration: 1. Assess the patient for need (clinical signs and symptoms, breath sounds, peak flow). 2. Gather the necessary equipment to deliver medication and select mask or mouthpiece delivery (nose clips may be needed with mouthpiece). 3. Wash or sanitize your hands and apply PPE. 4. Introduce yourself and your department. Verify patient identification and explain the procedure. 5. Position the patient in an upright seated position, if possible. 6. Aseptically fill the nebulizer with the ordered medication and diluent. Do not allow the tip of the saline vial to touch any inside surface of the nebulizer! 7. Set the compressed gas source to 6 to 10 L/m. 8. Coach the patient to breathe slowly through the mouth at normal VT. 9. Periodically reassess pulse throughout the treatment. Modify the patient’s technique as needed based on response and reinstruct as necessary. 10. Terminate the treatment when the complete medication dosage is nebulized or significant adverse reactions occur. 11. Reassess vital signs, breath sounds, and peak flow. And monitor the patient for adverse responses. 12. Rinse the nebulizer with sterile water and air dry or discard it between treatments. NOTE: The nebulizer should not be rinsed with tap water because of the possible contamination with Legionella. 34  Humidity and O2 Therapy The major functions of the upper airway are to warm, filter, and humidify the air that is breathed. Many conditions, such as the presence of an artificial airway, dehydration, fever, and the breathing of anhydrous gases, alter the efficiency of the upper airway in performing these functions. In these circumstances, respiratory therapist may deliver adjunctive humidity therapy to patients to minimize mucosal drying and irritation and to prevent secretions from becoming inspissated. Bubble humidifier: A bubble humidifier breaks (diffuse) an underwater gas stream into small bubbles. 1. Fill the humidifier to the fill line with sterile water. 2. Connect the bubble humidifier to the DISS outlet of a flowmeter. Adjust the flow rate to 5 Lpm. 35 Heat and moisture exchangers: Heat and moisture exchange (HME) units typically comprise a passive humidifier, also described as an “artificial nose.” Similar to the nose, such a device captures exhaled heat and moisture and returns up to 70% of the heat and humidity to the patient during the next inspiration. The three basic types of HME units are (1) simple condenser humidifiers, (2) hygroscopic condenser humidifiers, and (3) hydrophobic condenser humidifiers. 36 37 Oxygen (O2) Therapy: Oxygen is one of the most commonly administered respiratory drugs. Although administration of oxygen requires relatively simple skills, it is often misused or abused. The primary goal of oxygen therapy is to reduce morbidity and mortality associated with hypoxia. Proper administration, using the correct device, and assessing the adequacy of the delivered FIO2 can have a significant positive impact on patient outcomes. Nasal Cannula: Low flow rate of oxygen Used mostly for COPD patients (for long-term use) Flow rate of 1- 6 L/min Oxygen concentration of 24% -40% Figure 3 Nasal Cannula Simple Face Mask: Flow rate: 5 - 10 L/ min Oxygen concentration: 35% - 50% Minimum flow rate is 5 L/min 38 Figure 4 Simple face mask Partial Rebreather mask Mask with Reservoir beg which allows 1/3 of exhaled air to breathe. Flow rate minimum of 10 L/ min. Oxygen concentration: 40 -70% Non-Rebreather mask It is a plastic mask with a Reservoir bag. Consist of the one-way valve which prevents room air & exhaled air from entering the bag. Oxygen method of highest percentage oxygen concentration delivery system. Flow rate 10-15 L / minute Oxygen concentration: 95 - 100 % 39 Venturi Mask: Most accurate (fixed) oxygen concentration delivery system Flow rate 2 – 15 L/ min Oxygen concentration 24 – 60% Use acute respiratory distress. 40 41  Chest Physiotherapy Increased mucus production or impaired cough ability from physical limitations can overwhelm the body’s normal clearance mechanisms and lead to retained secretions. Removal of retained secretions and improved cough effort are necessary to prevent atelectasis and infection. Chest physiotherapy (CPT) includes: A. Postural drainage B. Chest percussion C. Expiratory vibration Postural drainage positions and procedure: Postural drainage is specific positions that allow the force of gravity to assist in the removal of bronchial secretions. The secretions drain from the affected bronchioles into the bronchi and trachea and are removed by coughing or Suctioning. Equipment: gloves, stethoscope, towel, pillow, electric bed, sputum container. Procedure: 1. Hand washing and wear gloves 2. Coordinate therapy 2 hours before meals and tube feedings. 3. Introduce yourself to the “patient” and explain the procedure. 4. Assess the patient before therapy. Assessment should include the following: pulse rate, respiratory rate, Spo2, blood pressure, level of dyspnea, and level of consciousness. 5. Auscultate the lung field before the procedure to identify the areas that need to be drained. 6. Place the patient in the appropriate positions for drainage depending on the affected lobes/ segments (Figure 1). Begin with from most-dependent to least-dependent lobes and segments (in adults, this is from bases to apices). 7. Use pillows and foam wedges to help maintain the proper positions and ensure patient comfort. 8. Positions are held for 3 - 15 minutes and modified according to the patient's condition and tolerance. 9. Monitored before, during, and after: subjective responses (pain, discomfort, dyspnea, response to therapy), breathing pattern, and sputum production. 10. Provide O2 supplement if SpO2 decreased during the session 42 Figure 1 43 The thick secretions that are difficult to cough up may be loosened by tapping (percussing) and vibrating the chest. Chest percussion and vibration help to dislodge mucus adhering to the bronchioles and bronchi. Manual percussion technique: 1. Remove your jewelry, wash your hands, and apply BSI. 2. Cup your hands and allow for relaxed motion from the wrist. Do not stiffen your upper arms. 3. Rhythmically strike the designated area, alternating hands. 4. Position the patient for postural drainage (any position may be used for this exercise). 5. Perform chest percussion on the patient for at least 3 minutes. Mechanical percussion devices can also be used with varying speed and force. Remember the following precautions: A. Do not percuss over any buttons, zippers, or similar items that the subject may have on. B. Do not percuss on bare skin. On an actual patient, use a towel or sheet over the patient’s skin. C. Avoid bony processes. Do not percuss on the spine, on clavicles, on scapulae, on breast tissue, over areas of incisions, over areas of rib fractures, or below the rib margins. Mechanical percussor Percussion cups 44 Expiratory vibration: Apply a gentle vibrating motion during exhalation only. Your upper arm muscles should tighten and allow transmission of the vibration to your hands. Do not shake the patient. Repeat the vibration technique two or three times for each segment. Expiratory vibration technique 45 References I. EGAN’S FUNDAMENTALS OF RESPIRATORY CARE, 12th Edition 2021 II. Basic Clinical Lab Competencies for Respiratory Care: An Integrated Approach, Fifth Edition Gary C. White 2013 III. Wilkins' Clinical Assessment in Respiratory Care – 9th edition Evolve - ELSEVIER ISBN: 9780323697019 2022 IV. Laboratory Exercises for Competency in Respiratory Care, Thomas J. Butler, PhD, RRT-NPS, RPFT, 3rd Edition 2013 V. American Association for Respiratory Care. (2020). AARC clinical practice guideline: Management of airway emergencies. Respiratory Care, 40(7), 749–760. American Association for Respiratory Care. (2019). AARC clinical practice guideline: Resuscitation and defibrillation in the health care setting. Respiratory Care, 49, 1085–1099. American Heart Association. (2018). Adjuncts for airway control and ventilation. Circulation, 112(11), IV-51–IV-57. MacIntyre, N. (2019). Discontinuing mechanical ventilatory support. Chest, 132(3), 1049–1056. Mokhlesi, B. (2018). Predicting extubation failure after successful completion of a spontaneous breathing trial. Respiratory Care, 52(12), 1710–1717. 46