- Email: [email protected]
Clinical Issues—May 2015
CONTINUING EDUCATION
Clinical Issues
2.6
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AMBER WOOD, MSN, RN, CNOR, CIC; SHARON A. VAN WICKLIN, MSN, RN, CNOR, CRNFA, CPSN-R, PLNC; SCOTT A. BRUBAKER, CTBS; LISA SPRUCE, DNP, RN, CNS-CP, ACNS, ACNP, ANP, CNOR Continuing Education Contact Hours
Approvals
indicates that continuing education (CE) contact hours are available for this activity. Earn the CE contact hours by reading this article, reviewing the purpose/goal and objectives, and completing the online Learner Evaluation at http://www .aorn.org/CE. Each applicant who successfully completes this program can immediately print a certificate of completion.
This program meets criteria for CNOR and CRNFA recertification, as well as other CE requirements.
Event: #15516 Session: #0001 Fee: Members $20.80, Nonmembers $41.60 The contact hours for this article expire May 31, 2018. Pricing is subject to change.
Purpose/Goal To provide the learner with knowledge of standard Ebola precautions related to personal protective equipment as well as AORN’s guidelines related to contaminated rigid sterilization containers, requirements for storing autologous tissue, storing autologous human vein autografts, dropped autografts, and antimicrobial and fluid barrier fabrics.
Objectives 1. Discuss practices that could jeopardize safety in the perioperative area. 2. Discuss common areas of concern that relate to perioperative best practices. 3. Describe implementation of evidence-based practice in relation to perioperative nursing care.
AORN is provider-approved by the California Board of Registered Nursing, Provider Number CEP 13019. Check with your state board of nursing for acceptance of this activity for relicensure.
Conflict-of-Interest Disclosures Sharon A. Van Wicklin, MSN, RN, CNOR, CRNFA, CPSN-R, PLNC; Scott A. Brubaker, CTBS; Lisa Spruce, DNP, RN, CNS-CP, ACNS, ACNP, ANP, CNOR; and Amber Wood, MSN, RN, CNOR, CIC, have no declared affiliations that could be perceived as posing potential conflicts of interest in the publication of this article. The behavioral objectives for this program were created by Helen Starbuck Pashley, MA, BSN, CNOR, clinical editor, with consultation from Susan Bakewell, MS, RN-BC, director, Perioperative Education. Ms Starbuck Pashley and Ms Bakewell have no declared affiliations that could be perceived as posing potential conflicts of interest in the publication of this article.
Sponsorship or Commercial Support No sponsorship or commercial support was received for this article.
Disclaimer Accreditation AORN is accredited as a provider of continuing nursing education by the American Nurses Credentialing Center’s Commission on Accreditation.
AORN recognizes these activities as CE for RNs. This recognition does not imply that AORN or the American Nurses Credentialing Center approves or endorses products mentioned in the activity. http://dx.doi.org/10.1016/j.aorn.2015.02.010 ª AORN, Inc, 2015
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CLINICAL ISSUES
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THIS MONTH Update on perioperative Ebola precautions Key words: Ebola, PPE, airborne, droplet, respirator. Contaminated rigid sterilization containers Key words: rigid sterilization containers, filters, valves, latches, gaskets. Requirements for storing autologous tissue Key words: tissue bank, autologous tissue, US Food and Drug Administration, human cells, delayed replantation or autotransplantation. Storing autologous human vein autografts Key words: autografts, storage media, Tiprotec, X-Vivo 10, American Association of Tissue Banks. Dropped autografts Key words: autograft, pulse lavage, wound classification, dropped grafts, cranial bone graft. Antimicrobial and fluid barrier fabrics Key words: antimicrobial fabrics, fluid barrier fabrics, surgical site infections, scrub attire, surgical attire.
Update on perioperative Ebola precautions QUESTION:
ANSWER:
Have there been any changes to personal protective equipment (PPE) recommendations when caring for a patient with known or suspected Ebola?
Perioperative team members should continue to take standard, contact, droplet, and airborne precautions when caring for a patient with known or suspected Ebola in the OR. The
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Table 1. PPE for Facilities to Follow When Caring for a Patient With Ebola
1,2
Nonscrubbed team members (eg, RN circulator, anesthesia professional) Follow PPE for surgical N95 respiratora Gown: fluid-resistant, extends to midcalf or Coveralls: fluid-resistant, without integrated hood Boot covers: fluid-resistant, extend to midcalf Gloves: double; outer pair has extended cuff Apron: fluid-resistant, extends to midcalf Full face shield and surgical hood No exposed skin!
Use of PAPRs based on risk assessment: Follow PPE for NIOSH-certified PAPRs Gown: fluid-resistant, extends to midcalf or Coveralls: fluid-resistant, without integrated hood Boot covers: fluid-resistant, extend to midcalf Gloves: double; outer pair has extended cuff Apron: fluid-resistant, extends to midcalf PAPR integrated with full face shield and hood No exposed skin! Scrubbed team members
a
Follow PPE for surgical N95 respirator Sterile surgical gown for AAMI’s level 4 criteria3 Boot covers: fluid-resistant, extend to midcalf Gloves: double, sterile Apron: fluid-resistant, extends to midcalf under sterile gown Full face shield and surgical hood or Surgical helmet system Full face shield and surgical hood or Surgical helmet system No exposed skin!
Do not follow PPE for PAPRs PAPRs should not be used where there is a sterile field during an operative or other invasive procedure.4 Follow PPE for surgical N95 respirator.
Sterile processing team members Option 1: Follow PPE for surgical N95 respiratora Gown5: fluid-resistant, extends to midcalf or Coveralls: fluid-resistant, without integrated hood Boot covers5: fluid-resistant, extend to midcalf Gloves: double; outer pair are general-purpose utility gloves with a cuff that extends beyond the cuff of the gown5 Apron: fluid-resistant, extends to midcalf Full face shield and surgical hood No exposed skin!
Option 2: Follow PPE for NIOSH-certified PAPRs Gown5: fluid-resistant, extends to midcalf or Coveralls: fluid-resistant, without integrated hood Boot covers5: fluid-resistant, extend to midcalf Gloves: double; outer pair are general-purpose utility gloves with a cuff that extends beyond the cuff of the gown5 Apron: fluid-resistant, extends to midcalf PAPR integrated with full face shield and hood No exposed skin!
PPE ¼ personal protective equipment; PAPRs ¼ powered air purifying respirators; NIOSH ¼ National Institute for Occupational Safety and Health; AAMI ¼ Association for the Advancement of Medical Instrumentation. a NIOSH-approved N95 respirator that has also been cleared by the US Food and Drug Administration. References 1. Infection prevention and control recommendations for hospitalized patients with known or suspected Ebola virus disease in U.S. hospitals. Centers for Disease Control and Prevention. http://www.cdc.gov/vhf/ebola/hcp/infection-prevention-and-control-recommendations.html. Updated January 9, 2015. Accessed January 16, 2015. 2. Guidance on personal protective equipment to be used by healthcare workers during management of patients with Ebola virus disease in U.S. hospitals, including procedures for putting on (donning) and removing (doffing). Centers for Disease Control and Prevention. http://www.cdc .gov/vhf/ebola/hcp/procedures-for-ppe.html. Updated October 20, 2014. Accessed January 16, 2015. 3. Guideline for sterile technique. In: Guidelines for Perioperative Practice. Denver, CO: AORN, Inc; 2015:67-96. 4. NPPTL Respirator Trusted-Source Information Section 1: NIOSH-Approved Respirators e What are they? How can they be identified? Where can I get them? http://www.cdc.gov/niosh/npptl/topics/respirators/ disp_part/respsource1.html. Accessed January 16, 2015. 5. Guideline for cleaning and care of surgical instruments and powered equipment. In: Guidelines for Perioperative Practice. Denver, CO: AORN, Inc; 2015:615-650.
Centers for Disease Control and Prevention (CDC) issued updated personal protective equipment (PPE) requirements in the Infection Prevention and Control Recommendations for 576 j AORN Journal
Hospitalized Patients With Known or Suspected Ebola Virus Disease in U.S. Hospitals.1 According to the CDC, when following facility requirements for PPE, perioperative
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Clinical Issues
personnel should have no exposed skin.2 See Table 1 for PPE recommendations in the perioperative setting.
http://www.cdc.gov/vhf/ebola/hcp/procedures-for-ppe.html. Updated October 20, 2014. Accessed January 16, 2015.
References
Amber Wood, MSN, RN, CNOR, CIC
1. Infection prevention and control recommendations for hospitalized patients with known or suspected Ebola virus disease in U.S. hospitals. Centers for Disease Control and Prevention. http://www.cdc .gov/vhf/ebola/hcp/infection-prevention-and-control-recommendations .html. Updated January 9, 2015. Accessed January 16, 2015. 2. Guidance on personal protective equipment to be used by healthcare workers during management of patients with Ebola virus disease in U.S. hospitals, including procedures for putting on (donning) and removing (doffing). Centers for Disease Control and Prevention.
Perioperative Nursing Specialist AORN Center for Nursing Practice
Contaminated rigid sterilization containers QUESTION: I heard that a European study showed bacterial growth on rigid sterilization containers after sterilization. If so, does this present a risk of infection for the patient?
ANSWER: Dunkelberg and Fleitmann-Glende1 examined the microbial growth and barrier properties of 216 sterilization containers after sterilization in a laboratory environment. The containers were obtained from central sterile supply departments from four different hospitals in Germany. The researchers theorized that the integrity of sterilization containers could be affected by repeated use and sterilization; the seal of the container could be weakened by exposure to heat and mechanical stress, causing rivets, bolts, and nuts to loosen or the latching mechanism that secures the lid to the base to deteriorate; and a risk of contamination from an exchange of air between the lower pressure inside of the container and the higher pressure in the environment could result, as might occur when the container is transferred to an area with positive pressure such as the OR. The containers used in the study included containers with paper filters, reusable textile filters, and permanent plastic filters. The researchers placed uncovered thermoresistant plates filled with Sabouraud agar in the base of the containers before sterilizing them. To determine the microbial barrier function of the containers, they exposed the containers to an aerosol suspension of Saccharomyces cerevisiae after sterilization. To determine the potential for microbial contamination caused by changes in atmospheric pressure, the researchers exposed 12 containers to 24 different pressure changes followed by a microbial aerosol of S cerevisiae.
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The researchers found that only two of 11 containers with paper filters and nine of 79 containers with textile filters lacked microbial growth; however, 14 of 15 containers with permanent plastic filters showed no growth. Notably, the contaminated containers showed visible defects, including fissured or loosened lid seals. The researchers found that only four of the 12 containers exposed to the pressure changes maintained a high microbial barrier efficacy. The researchers demonstrated the need for challenging the effectiveness of the microbial barrier of rigid sterilization containers and a method to do so. Although the generalizability of this study may be limited because the containers used by the researchers may differ from those used in the United States, the results of the study emphasize the importance of a quality assurance program that includes careful and periodic inspections of the containers before use in a manner according to the manufacturer’s written instructions for use.2 Personnel should inspect containers to ensure the valves work freely and the sealing surfaces and edges are aligned correctly and are free from dents or chips3; check gaskets to ensure they are pliable, securely fastened, and without breaks or cuts3; inspect filter retention mechanisms and fasteners (eg, screws, rivets) to verify they are secure and functioning correctly; check filter materials for integrity and freedom from holes3; and repair or replace defective containers. The researchers emphasized that although there was no evidence that failure of a sterilization container had caused patient infection,
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the results of their study strongly support the need for monitoring the integrity of packaging systems at least annually.
Sharon A. Van Wicklin, MSN, RN, CNOR, CRNFA, CPSN-R, PLNC
References 1. Dunkelberg H, Fleitmann-Glende F. Measurement of the microbial barrier effectiveness of sterilization containers in terms of the log reduction value for prevention of nosocomial infections. Am J Infect Control. 2006;34(5):285-289. 2. Guideline for sterilization. In: Guidelines for Perioperative Practice. Denver, CO: AORN, Inc; 2015:665-692. 3. Association for the Advancement of Medical Instrumentation (AAMI). ANSI/AAMI ST79:2010/ A4:2013: Comprehensive Guide to Steam Sterilization and Sterility Assurance in Health Care Facilities. Arlington, VA: AAMI; 2013.
Perioperative Nursing Specialist AORN Nursing Department
Requirements for storing autologous tissue QUESTION: Is it true that if a facility stores autologous tissue, the facility must be registered with the US Food and Drug Administration (FDA) as a tissue bank?
ANSWER: A facility that handles autologous tissue for delayed replantation or autotransplantation within the same facility is not required to register with the US Food and Drug Administration (FDA) as a tissue establishment (ie, tissue bank) and is not required to follow the requirements of the FDA regulations for cellular and tissue-based products (x21 CFR Part 1271).1,2 Facilities or health care organizations that handle autologous tissue are required to recover, process, package, label, store, track, and replant or autotransplant the tissue in a manner that minimizes microbial growth, prevents replantation or autotransplantation to the wrong patient, and reduces the risk for errors.1 Although the regulation defines manufacturing to include recovery, processing, storage, labeling, packaging, or distribution of any human cell or tissue,1,3 the FDA considers most procedures related to autologous tissue to be a single procedure encompassed within the element of storage. The reader can refer to section 1271.15(b), of the regulation, in which storage of autologous tissue is exempt if replantation or autotransplantation will occur in the facility where the recovery took place.2 Similarly, packaging and labeling of the autologous tissue can be encompassed within the exception for storage.2 Freezing autologous tissue as a method of storage does not, in
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itself, require the facility to register with the FDA as a tissue establishment.2 In addition, the FDA has interpreted the “same surgical procedure” language in its final rule to include recovery and storage before replantation or autotransplantation.1 Retaining autologous tissue to be used in a subsequent application for the same patient is exempt from registration because the two applications are essentially a single, continuous procedure.2 The facility is required to register with the FDA if autologous tissue handling includes steps to process the autograft that require specific manufacturing controls to decontaminate the tissue (eg, subjecting the autograft to a steam sterilization process).2 If autologous tissueehandling functions are expanded to include distribution of the autograft to another facility located at a different address, registration and listing with the FDA using Form FDA 3356 is required.1
References
1. x21 CFR 1271: Human cells, tissues, and cellular and tissue-based products. April; 2013. US Food and Drug Administration. http:// www.accessdata.fda.gov/scripts/cdrh/cfdocs/cfcfr/CFRSearch .cfm?CFRPart¼1271. Accessed December 6, 2014. 2. Human Cells, Tissues, and Cellular and Tissue-Based Products; Establishment Registration and Listing. Fed Regist. 2001;66(13): 5447-5469. http://www.gpo.gov/fdsys/pkg/FR-2001-01-19/pdf/ 01-1126.pdf. Accessed December 6, 2014. 3. Current Good Tissue Practice (CGTP) and Additional Requirements for Manufacturers of Human Cells, Tissues, and Cellular and TissueBased Products (HCT/ Ps). Silver Spring, MD: US Food and Drug Administration; 2012.
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Clinical Issues
Sharon A. Van Wicklin, MSN, RN, CNOR, CRNFA, CPSN-R, PLNC
Scott A. Brubaker, CTBS Chief Policy Officer American Association of Tissue Banks McLean, VA
Perioperative Nursing Specialist AORN Nursing Department
Storing autologous human vein autografts QUESTION: How should autologous human vein autografts be stored?
ANSWER: The American Association of Tissue Banks recommends that autologous vein autografts be stored between 32 F to 50 F (0 C to 10 C) for no longer than 14 days.1 Refrigerated veins should be submerged in storage medium, and the storage medium should be changed every 72 hours using sterile technique.2 Keeping the veins submerged helps prevent dehydration of the tissue.2 Changing the storage medium every 72 hours may help prevent microbial growth and improve vein autograft viability.3,4 Three studies provide moderate evidence that regularly changing the storage medium of human vein autografts may extend viability.5-7 To investigate the effect of storing saphenous vein segments in normal saline solution compared with TiProtec solution, Wilbring et al5 isolated 19 saphenous vein segments from patients undergoing coronary artery bypass grafting with autologous saphenous veins in a university heart center between October 2008 and March 2010. The veins were extracted without the use of electrocoagulation. Each vein segment was divided into two parts and placed in either normal saline solution or TiProtec. The vein segments were stored at 39.2 F (4 C) for 96 hours and then examined. The researchers found that the vessel function was significantly reduced after 24 hours of cold storage in normal saline solution. After 96 hours of storage in normal saline solution, there was minimal to no vessel function remaining. Vascular function of the vein segments stored in TiProtec was significantly better preserved. The researchers concluded that cold storage of venous grafts was feasible for as long as 96 hours when the grafts were
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stored in TiProtec solution; however, storage of venous grafts in normal saline solution was not recommended beyond 24 hours.5 Molnar et al6 conducted a quasi-experimental study to investigate the passive and active biomechanical properties of 72 saphenous vein segments remaining after coronary bypass grafting from 32 patients in a university medical center. The vein segments were divided into eight testing groups: fresh, examined immediately after recovery; stored in normal Krebs-Ringer solution at 32 F to 39.2 F (0 C to 4 C), examined one week after recovery; stored in normal Krebs-Ringer solution at 32 F to 39.2 F (0 C to 4 C), examined two weeks after recovery; stored in X-Vivo 10TM at 32 F to 39.2 F (0 C to 4 C), examined one week after recovery; stored in X-Vivo 10 at 32 F to 39.2 F (0 C to 4 C), examined two weeks after recovery; stored in X-Vivo 10 at 32 F to 39.2 F (0 C to 4 C), examined three weeks after recovery; stored in X-Vivo 10 at 32 F to 39.2 F (0 C to 4 C), examined four weeks after recovery; or cryopreserved at -220 F (-140 C), examined three weeks after recovery. Researchers thawed the cryopreserved samples by immersion in warm (98.6 F [37 C]) Krebs-Ringer solution. Biomechanical testing showed that the vein segments stored in Krebs-Ringer solution lost their ability to dilate and contract within one week. The segments stored in X-Vivo 10 preserved their contractility after one week, and contractility slowly decreased during the fourweek study period. There was a slight decrease in vessel wall thickness, but the lumen diameter was not affected. The elastic parameters were almost identical to the fresh segment; however, AORN Journal j 579
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the cryopreserved segments narrowed, the vessel wall thickened, and contractility diminished. The researchers concluded that storage in X-Vivo 10 helped to preserve the passive and active biomechanical properties of human saphenous vein segments. They also concluded that maintaining these properties could be expected to improve stored autologous vein graft viability.6 Baumann et al7 used light transmission and scanning electron microscopy to investigate the effects of various methods of vein preparation on endothelial and smooth muscle cells in cephalic veins from dogs. The researchers removed the veins, divided them into 10-cm segments (N ¼ 10), placed them in one of three solutions (ie, autologous blood [n ¼ 3], Plasma-Lyte [n ¼ 2], or Plasma-Lyte with 0.6 mg/mL papaverine [n ¼ 5]) containing heparin 10,000 units/L, and then stored them at 50 F (10 C) for either five minutes or one hour. The researchers found the vein wall was extremely sensitive to dissection, manipulation, or introduction of fixative solutions and reacted to such stimulations with severe contraction that not only diminished the luminal diameter but also resulted in the protrusion of endothelial cells into the lumen and the formation of cytoplasmic extensions of medial smooth muscle cells. The veins stored in autologous blood demonstrated the greatest amount of vessel wall contraction and endothelial cell loss. Veins stored in Plasma-Lyte for five minutes showed few contractions; however, after one hour, there were contractions and some endothelial cell loss. Veins stored in Plasma-Lyte supplemented with papaverine had the most relaxed appearance and minimal endothelial cell loss. The researchers recommended mitigating damaging vein graft contractions by using gentle surgical dissection to avoid vein spasm; instituting and intermittently repeating (ie, every two to three minutes) gentle perfusion with a solution of Plasma-Lyte supplemented with papaverine immediately after exposure of the distal end of the vein and while the rest of the vein is removed; avoiding the use of autologous blood for vein immersion, storage, or distention; and monitoring the pressure used to check vein grafts for leaks at less than 100 mmHg. The researchers concluded that attention to the details of vein dissection, preparation, and storage medium could lead to a significant improvement in endothelial preservation and subsequent patency rates after cardiac bypass procedures.7 There are gaps in the literature related to best practices for storage of autologous veins, and further research is warranted.
the Lonza Group, Walkersville, MD. Plasma-Lyte is a registered trademark of Baxter Healthcare, Deerfield, IL.
References 1. Standards for Tissue Banking. McLean, VA: American Association of Tissue Banks; 2012. 2. Guideline for autologous tissue management. In: Guidelines for Perioperative Practice. Denver, CO: AORN, Inc; 2015:187-238. 3. Robb EC, Bechmann N, Plessinger RT, Boyce ST, Warden GD, Kagan RJ. Storage media and temperature maintain normal anatomy of cadaveric human skin for transplantation to full-thickness skin wounds. J Burn Care Rehabil. 2001;22(6):393-396. 4. Cram AE, Domayer MA. Short-term preservation of human autografts. J Trauma. 1983;23(10):872-873. 5. Wilbring M, Tugtekin SM, Zatschler B, et al. Preservation of endothelial vascular function of saphenous vein grafts after long-time storage with a recently developed potassium-chloride and N-acetylhistidine enriched storage solution. Thorac Cardiovasc Surg. 2013;61(8):656-662. 6. Molnar GF, Nemes A, Kekesi V, Monos E, Nadasy GL. Maintained geometry, elasticity and contractility of human saphenous vein segments stored in a complex tissue culture medium. Eur J Vasc Endovasc Surg. 2010;40(1):88-93. 7. Baumann FG, Catinella FP, Cunningham JN Jr, Spencer FC. Vein contraction and smooth muscle cell extensions as causes of endothelial damage during graft preparation. Ann Surg. 1981;194(2):199-211.
Sharon A. Van Wicklin, MSN, RN, CNOR, CRNFA, CPSN-R, PLNC Perioperative Nursing Specialist AORN Nursing Department
Scott A. Brubaker, CTBS Chief Policy Officer American Association of Tissue Banks McLean, VA
Editor’s notes: TiProtec is a registered trademark of Dr Franz K€ohler Chemie, Bensheim, Germany. X-Vivo 10 is a trademark of
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Dropped autografts QUESTION: Recently, one of our surgeons dropped the autologous cranial bone flap during the replantation procedure and then replanted the contaminated autograft. How should this event have been managed?
ANSWER: The collective evidence indicates there are instances in which autologous grafts necessary for replantation or autotransplantation have been contaminated or dropped on the OR floor and that the infection rate associated with replantation or autotransplantation of contaminated grafts is low.1-3 Various processes for decontaminating the graft have been implemented; however, there is no consistent protocol for managing the event.1-3 Further research is warranted on best practices for decontaminating contaminated autologous grafts and determining methods to prevent the graft from being dropped or contaminated. If an autologous graft intended for replantation or autotransplantation is dropped or contaminated during the procedure, a multidisciplinary team consisting of the surgeon, perioperative RN, and infection preventionist should conduct a risk assessment to consider the benefits and potential harms associated with replantation or autotransplantation of the contaminated autograft compared with other treatment options (eg, discarding the graft and using artificial material).4 If a decision is made to replant or autotransplant the contaminated autograft, the following steps should be taken: Rinse the contaminated graft in sterile normal saline solution to remove surface debris and contaminants. Rinsing the dropped autograft in normal saline solution may be sufficient to remove surface debris and contaminants.4 In a randomized controlled trial conducted to determine the amount of contamination that occurred when a bone graft was dropped on the floor, Presnal and Kimbrough5 concluded that replanting a dropped bone graft was acceptable without extensive, and potentially damaging, efforts to disinfect the graft.5 Use pulsatile lavage at low-pressure settings (eg, 6 psi to 14 psi)6,7 and use sterile normal saline solution for more thorough cleansing of contaminated bone grafts if indicated (eg, adherent debris is present).4 Pulsatile lavage may be useful for removing soil or microorganisms that are difficult to kill or remove with rinsing, such as heavy contamination and debris embedded in the tissue (eg, from a traumatic accident). There is no agreement in the literature as to the absolute definition of
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low-pressure versus high-pressure lavage.6 Bhandari et al8 found that high-pressure lavage (ie, 70 psi) may damage the bone and carry surface contaminants deeper into the bone. In another study, Bhandari et al6 found that low-pressure lavage (ie, 14 psi) was as effective as high-pressure lavage in removing adherent bacteria from bone and preserved the bony architecture when applied within three hours of contamination. Use a separate sterile field for decontaminating the dropped graft and exercise care to prevent splashing onto the primary sterile field.4 Using a separate sterile field may help prevent contamination of the primary sterile field by microorganisms on the contaminated autograft and from droplets caused by rinsing or irrigation fluid used to decontaminate the graft.4 Implementing corrective actions to maintain the sterility of the surgical field may prevent or reduce microbial contamination and help minimize the patient’s risk of surgical site infection.4 Implement corrective actions as necessary to maintain the sterility of the primary sterile field (eg, changing gowns and gloves after pulsatile lavage of the contaminated graft).4 Preventing or reducing microbial contamination helps minimize the patient’s risk of surgical site infection.4 Change the wound classification to Class III, Contaminated.4,9 Replantation or autotransplantation of a contaminated autograft constitutes a major break in sterile technique. According to the Centers for Disease Control and Prevention surgical wound classification system, a surgical wound with a major break in sterile technique is classified as Class III, Contaminated.9 Document the event in a variance report.4 Variance reports document the steps taken during the event and provide a mechanism for alerting infection preventionists to the need for surveillance.4 Variance reports also can be useful for quality improvement activities.4 Conduct a debriefing session and a root cause analysis with members of the surgical team and other individuals who may be helpful in providing a critical analysis and determining the factors that contributed to the event and methods to prevent its recurrence.4 Implementing a process of debriefing and root cause analysis after the event may help prevent future incidents by examining underlying factors and system flaws that may have contributed to the event and may respond to analysis and correction.10
In addition, the following actions may be taken: Add antibiotics or antiseptics to the solution used to rinse the autograft.4 Adding antibiotics or antiseptics to the irrigation solution may be unnecessary or potentially harmful. Antibiotics
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and antiseptics may not have been validated for use in irrigation solutions and may therefore pose a risk to the patient. Tissue proteins may inhibit the effectiveness of the antibiotics.11 Some antibiotics, such as bacitracin,12 have not been found to be effective for eliminating bone contaminants, whereas others, such as rifamycin,13 have been found to be effective for decontaminating contaminated bone grafts. Antiseptic solutions are intended for external use and may be ineffective or toxic when used internally. A povidone-iodine solution has been found to provide effective bone decontamination14; however, the antibacterial activity of 10% povidone-iodine solution is directly related to the level of contamination and the duration of exposure to the contaminated bone.15 Lacey16 found that the antibacterial effect of a povidone-iodine solution was inactivated in the presence of blood. Kaysinger et al17 found that 0.5%, 5%, and 50% povidone-iodine solutions and 1.5% and 3% hydrogen peroxide (H2O2) solutions were cytotoxic to osteoblasts. Bhandari et al6 found that a povidone-iodine solution and bacitracin decreased the number of osteoclasts and impaired osteoblast function. When used to disinfect explanted bone, Sch€ ultke et al18 found that H2O2 was ineffective. Studies have shown chlorhexidine gluconate to be both effective14 and ineffective13 for decontaminating contaminated bone grafts and it has also been found to be toxic to bone cells, even at very low concentrations (ie, 1%).12 Using chemical disinfectants (eg, glutaraldehyde) on bone may cause tissue damage or the disinfectants may be difficult to remove.4 Take samples for culture before and after decontamination of the graft to determine the identity of the contaminating microorganism and the level of contamination.4 Tissue cultures may be helpful in determining whether the dropped autograft was contaminated before or after decontamination measures were implemented.4 The results of the cultures may be useful in guiding treatment after replantation.4 Consult with an infection preventionist to assess the benefits versus harms of implementing postoperative broad-spectrum antibiotic prophylaxis therapy.4 Send the autograft to a tissue bank for decontamination and processing.4 Some tissue banks provide services to decontaminate autologous bone skull flaps when contamination is suspected (ie, dropped autograft, head trauma, positive culture result).4 The contaminated graft should not be subjected to the steam sterilization process,4 which denatures bone proteins, may severely damage the bone structure, and increases the potential for bone resorption and infection.18 The steam sterilization process has not been validated for use with human tissue. A significantly high rate of postoperative graft infections has been found when cranial bone grafts subjected to the steam sterilization process were used for cranial reconstruction.19
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References
1. Jankowitz BT, Kondziolka DS. When the bone flap hits the floor. Neurosurgery. 2006;59(3):585-589. 2. Kang L, Mermel LA, Trafton PG. What happens when autogenous bone drops out of the sterile field during orthopaedic trauma surgery. J Orthop Trauma. 2008;22(6):430-431. 3. Centeno RF, Desai AR, Watson ME. Management of contaminated autologous grafts in plastic surgery. Eplasty. April 22, 2008;8:e23. 4. Guideline for autologous tissue management. In: Guidelines for Perioperative Practice. Denver, CO: AORN, Inc; 2015:187-238. 5. Presnal BP, Kimbrough EE. What to do about a dropped bone graft. Clin Orthop Relat Res. 1993;296:310-311. 6. Bhandari M, Schemitsch EH, Adili A, Lachowski RJ, Shaughnessy SG. High and low pressure pulsatile lavage of contaminated tibial fractures: an in vitro study of bacterial adherence and bone damage. J Orthop Trauma. 1999;13(8):526-533. 7. Hirn M, Laitinen M, Pirkkalainen S, Vuento R. Cefuroxime, rifampicin and pulse lavage in decontamination of allograft bone. J Hosp Infect. 2004;56(3):198-201. 8. Bhandari M, Adili A, Lachowski RJ. High pressure pulsatile lavage of contaminated human tibiae: an in vitro study. J Orthop Trauma. 1998;12(7):479-484. 9. Mangram AJ, Horan TC, Pearson ML, Silver LC, Jarvis WR. Guideline for Prevention of Surgical Site Infection, 1999. Centers for Disease Control and Prevention (CDC) Hospital Infection Control Practices Advisory Committee. Am J Infect Control. 1999;27(2): 97-132; quiz 133-134; discussion 96. 10. Makary MA, Holzmueller CG, Sexton JB, et al. Operating room debriefings. Jt Comm J Qual Patient Saf. 2006;32(7):407-410. 357. 11. Sample Procedure: Handling Autologous Bone Skull Flaps. Version 14. McLean, VA: American Association of Tissue Banks; 2012. 12. Bhandari M, Adili A, Schemitsch EH. The efficacy of low-pressure lavage with different irrigating solutions to remove adherent bacteria from bone. J Bone Joint Surg Am. 2001;83-A(3):412-419. 13. Yaman F, Unl€u G, Atilgan S, Celik Y, Ozekinci T, Yaldiz M. Microbiologic and histologic assessment of intentional bacterial contamination of bone grafts. J Oral Maxillofac Surg. 2007;65(8): 1490-1494. 14. Bruce B, Sheibani-Rad S, Appleyard D, et al. Are dropped osteoarticular bone fragments safely reimplantable in vivo? J Bone Joint Surg Am. 2011;93(5):430-438. 15. Soyer J, Rouil M, Castel O. The effect of 10% povidone-iodine solution on contaminated bone allografts. J Hosp Infect. 2002; 50(3):183-187. 16. Lacey RW. Antibacterial activity of povidone iodine towards nonsporing bacteria. J Appl Bacteriol. 1979;46(3):443-449. 17. Kaysinger KK, Nicholson NC, Ramp WK, Kellam JF. Toxic effects of wound irrigation solutions on cultured tibiae and osteoblasts. J Orthop Trauma. 1995;9(4):303-311. 18. Sch€ultke E, Hampl JA, Jatzwauk L, Krex D, Schackert G. An easy and safe method to store and disinfect explanted skull bone. Acta Neurochir (Wien). 1999;141(5):525-528. 19. Matsuno A, Tanaka H, Iwamuro H, et al. Analyses of the factors influencing bone graft infection after delayed cranioplasty. Acta Neurochir (Wien). 2006;148(5):535-540.
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May 2015, Volume 101, No. 5
Clinical Issues
Sharon A. Van Wicklin, MSN, RN, CNOR, CRNFA, CPSN-R, PLNC
Scott A. Brubaker, CTBS Chief Policy Officer American Association of Tissue Banks McLean, VA
Perioperative Nursing Specialist AORN Nursing Department
Antimicrobial and fluid barrier fabrics QUESTION: Are fabrics available that can prevent bacteria from being retained on the surface of the material? If so, can using these fabrics for scrub attire provide protection for perioperative personnel and also help prevent surgical site infections?
ANSWER: According to the AORN “Guideline for surgical attire,”1 “Scrub attire may be made of antimicrobial fabric.”1(p101) There is emerging evidence regarding the use of fabrics with fluid barrier and antimicrobial properties to prevent bacteria and fungi from adhering to the fabric.1 Bearman et al2 conducted a randomized controlled trial to determine the effectiveness of an antimicrobial, fluid barrier scrub fabric for reducing the bacterial burden on the hands of and scrub attire worn by health care workers in an intensive care unit setting of an academic medical center. All study participants (N ¼ 30) were randomly assigned to wear either traditional scrub attire or scrub attire made of antimicrobial, fluid barrier fabric during a clinical shift during a four-week period. The researchers took cultures from the abdominal and leg pockets of the scrub attire at the beginning and end of the shifts. Notably, the researchers did not specify the length of the clinical shifts. The researchers found a significant difference in the number of methicillin-resistant Staphylococcus aureus colony-forming units in both the leg and abdominal area of the antimicrobial, fluid barrier scrub attire compared with traditional scrub attire at the beginning and end of shifts. The researchers concluded that when combined with infection prevention strategies, such as hand hygiene and the use of personal protective equipment,
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the use of antimicrobial, fluid barrier scrub attire may limit the bacterial burden of the inanimate environment.2 In a quasi-experimental study to examine whether antibacterial finishes on fabric could effectively reduce the presence of bacteria on fabric used for health care workers’ uniforms, Chen-Yu et al3 found that the antibacterial finishes provided a significant reduction of S aureus and Klebsiella pneumoniae. The researchers concluded that adding antibacterial finishes to fabric was an effective method of reducing bacterial contamination. Mariscal et al4 conducted a quasi-experimental study to evaluate the action of a commercially available antimicrobial textile on 33 strains of bacteria. They found that the antimicrobial fabric significantly reduced the numbers of four reference microorganisms (ie, Escherichia coli, Pseudomonas aeruginosa, Morganella morgannii, S aureus) compared with the numbers of reference microorganisms on the control fabric. Sun et al5 quantitatively and qualitatively evaluated fabrics treated with antimicrobial chemicals in a nonexperimental study. The researchers found that even after 50 washings, the treated fabrics exhibited antibacterial properties against gram-negative (eg, Escherichia coli) and gram-positive (eg, S aureus) bacteria and fungi (eg, Candida albicans) within a two-minute contact time. Despite the biocidal properties of the antimicrobial fabric, the researchers found it was nontoxic to human skin. They concluded that antimicrobial fabrics were effective and suitable for medical use. In a randomized crossover trial conducted to determine whether applying an antimicrobial coating to scrub attire worn by nurses and patient care technicians working in the adult intensive and intermediate care units of a university medical center would reduce the overall rate of scrub contamination with pathogenic
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bacteria (eg, S aureus, Enterococcus species, gram-negative bacteria), Boutin et al6 cultured the front of 720 scrub tops worn by 90 health care workers during the last four hours of a 12-hour shift. The researchers found that 30% (n ¼ 217) of the scrub tops were contaminated with pathogenic bacteria. The treated scrub attire showed a contamination rate of 30% (n ¼ 108), and the untreated scrub attire showed a contamination rate of 30.3% (n ¼ 109). The researchers concluded the antimicrobial coating did not prevent bacterial contamination of scrub attire. In a quasi-experimental study to evaluate the bactericidal effects of woven and nonwoven fabrics coated with a hydroxyapatite-binding silver/titanium dioxide ceramic composite, Kasuga et al7 found that bacterial cell counts of S aureus and E coli on both the woven and nonwoven fabrics decreased to less than 2-log10 colony-forming units/mL within six hours and were undetectable at the end of the 18-hour incubation period. Bacterial cell counts of P aeruginosa could not be detected after three to six hours. In a nonexperimental study of silver-impregnated scrub attire, Gross et al8 compared the contamination rates of traditional ambulance service clothing and silver-impregnated ambulance service clothing worn by 10 emergency workers. The samples were taken from the right sleeves and front of the jackets and the right thighs of the pants before use and at the end of the third and seventh working days. Notably, the researchers did not specify the length of the working day. The researchers found no significant difference in the amount of contamination between the two clothing types; however, they recommended verifying the result of the study with a larger sample size.
5. Sun G, Qian L, Xu X. Antimicrobial and medical-use textiles. Textile Asia. 2001;32(9):33-35. 6. Boutin MA, Thom KA, Zhan M, Johnson JK. A randomized crossover trial to decrease bacterial contamination on hospital scrubs. Infect Control Hosp Epidemiol. 2014;35(11):1411-1413. 7. Kasuga E, Kawakami Y, Matsumoto T, et al. Bactericidal activities of woven cotton and nonwoven polypropylene fabrics coated with hydroxyapatite-binding silver/ titanium dioxide ceramic nanocomposite “Earth-plus.” Int J Nanomed. 2011;6:1937-1943. 8. Gross R, H€ubner N, Assadian O, Jibson B, Kramer A. Pilot study on the microbial contamination of conventional vs. silver-impregnated uniforms work by ambulance personnel during one week of emergency medical service. GMS Krankenhhyg Interdiszip. 2010; 5(2):1-16. 9. Bauer J, Kowal K, Tofail SAM, Podbielska H. MRSA-resistant textiles. In: Tofail SAM, ed. Biological Interactions with Surface Charge in Biomaterials. Cambridge, England: RSC Publishing; 2012:193-207. 10. Rajendran R, Radhai R, Kotresh TM, Csiszar E. Development of antimicrobial cotton fabrics using herb loaded nanoparticles. Carbohydr Polym. 2013;91(2):613-617.
Sharon A. Van Wicklin, MSN, RN, CNOR, CRNFA, CPSN-R, PLNC Perioperative Nursing Specialist AORN Nursing Department
Incorporating textile technologies into the material used for scrub attire and other garments worn by health care personnel may help prevent microorganisms from being acquired and retained on the fabric and reduce the risk of transmission of microorganisms that may cause surgical site infections.2-10 This issue warrants further research.
References 1. Guideline for surgical attire. In: Guidelines for Perioperative Practice. Denver, CO: AORN, Inc; 2015:97-120. 2. Bearman GM, Rosato A, Elam K, et al. A crossover trial of antimicrobial scrubs to reduce methicillin-resistant Staphylococcus aureus burden on healthcare worker apparel. Infect Control Hosp Epidemiol. 2012;33(3):268-275. 3. Chen-Yu JH, Eberhardt DM, Kincade DH. Antibacterial and laundering properties of AMS and PHMB as finishing agents on fabric for health care workers’ uniforms. Clothing Text Res J. 2007;25(3):258-272. 4. Mariscal A, Lopez-Gigosos RM, Carnero-Varo M, FernandezCrehuet J. Antimicrobial effect of medical textiles containing bioactive fibres. Eur J Clin Microbiol Infect Dis. 2011;30(2):227-232.
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Lisa Spruce, DNP, RN, CNS-CP, ACNS, ACNP, ANP, CNOR Director, Evidence-Based Perioperative Practice AORN Nursing Department
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LEARNER EVALUATION
Continuing Education: Clinical Issues 2.6
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T
his evaluation is used to determine the extent to which this continuing education program met your learning needs. The evaluation is printed here for your convenience. To receive continuing education credit, you must complete the online Learner Evaluation at http://www.aorn.org/CE. Rate the items as described below.
PURPOSE/GOAL To provide the learner with knowledge of standard Ebola precautions related to personal protective equipment as well as AORN’s guidelines related to contaminated rigid sterilization containers, requirements for storing autologous tissue, storing autologous human vein autografts, dropped autografts, and antimicrobial and fluid barrier fabrics.
6.
Will you be able to use the information from this article in your work setting? 1. Yes 2. No
7.
Will you change your practice as a result of reading this article? (If yes, answer question #7A. If no, answer question #7B.)
7A.
How will you change your practice? (Select all that apply) 1. I will provide education to my team regarding why change is needed. 2. I will work with management to change/implement a policy and procedure. 3. I will plan an informational meeting with physicians to seek their input and acceptance of the need for change. 4. I will implement change and evaluate the effect of the change at regular intervals until the change is incorporated as best practice. 5. Other: __________________________________
7B.
If you will not change your practice as a result of reading this article, why? (Select all that apply) 1. The content of the article is not relevant to my practice. 2. I do not have enough time to teach others about the purpose of the needed change. 3. I do not have management support to make a change. 4. Other: __________________________________
8.
Our accrediting body requires that we verify the time you needed to complete the 2.6 continuing education contact hour (156-minute) program: _________________________________
OBJECTIVES To what extent were the following objectives of this continuing education program achieved? 1. Discuss practices that could jeopardize safety in the perioperative area. Low 1. 2. 3. 4. 5. High 2.
Discuss common areas of concern that relate to perioperative best practices. Low 1. 2. 3. 4. 5. High
3.
Describe implementation of evidence-based practice in relation to perioperative nursing care. Low 1. 2. 3. 4. 5. High
CONTENT 4.
5.
To what extent did this article increase your knowledge of the subject matter? Low 1. 2. 3. 4. 5. High To what extent were your individual objectives met? Low 1. 2. 3. 4. 5. High
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AORN Journal j 585
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2.6
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AMBER WOOD, MSN, RN, CNOR, CIC; SHARON A. VAN WICKLIN, MSN, RN, CNOR, CRNFA, CPSN-R, PLNC; SCOTT A. BRUBAKER, CTBS; LISA SPRUCE, DNP, RN, CNS-CP, ACNS, ACNP, ANP, CNOR Continuing Education Contact Hours
Approvals
indicates that continuing education (CE) contact hours are available for this activity. Earn the CE contact hours by reading this article, reviewing the purpose/goal and objectives, and completing the online Learner Evaluation at http://www .aorn.org/CE. Each applicant who successfully completes this program can immediately print a certificate of completion.
This program meets criteria for CNOR and CRNFA recertification, as well as other CE requirements.
Event: #15516 Session: #0001 Fee: Members $20.80, Nonmembers $41.60 The contact hours for this article expire May 31, 2018. Pricing is subject to change.
Purpose/Goal To provide the learner with knowledge of standard Ebola precautions related to personal protective equipment as well as AORN’s guidelines related to contaminated rigid sterilization containers, requirements for storing autologous tissue, storing autologous human vein autografts, dropped autografts, and antimicrobial and fluid barrier fabrics.
Objectives 1. Discuss practices that could jeopardize safety in the perioperative area. 2. Discuss common areas of concern that relate to perioperative best practices. 3. Describe implementation of evidence-based practice in relation to perioperative nursing care.
AORN is provider-approved by the California Board of Registered Nursing, Provider Number CEP 13019. Check with your state board of nursing for acceptance of this activity for relicensure.
Conflict-of-Interest Disclosures Sharon A. Van Wicklin, MSN, RN, CNOR, CRNFA, CPSN-R, PLNC; Scott A. Brubaker, CTBS; Lisa Spruce, DNP, RN, CNS-CP, ACNS, ACNP, ANP, CNOR; and Amber Wood, MSN, RN, CNOR, CIC, have no declared affiliations that could be perceived as posing potential conflicts of interest in the publication of this article. The behavioral objectives for this program were created by Helen Starbuck Pashley, MA, BSN, CNOR, clinical editor, with consultation from Susan Bakewell, MS, RN-BC, director, Perioperative Education. Ms Starbuck Pashley and Ms Bakewell have no declared affiliations that could be perceived as posing potential conflicts of interest in the publication of this article.
Sponsorship or Commercial Support No sponsorship or commercial support was received for this article.
Disclaimer Accreditation AORN is accredited as a provider of continuing nursing education by the American Nurses Credentialing Center’s Commission on Accreditation.
AORN recognizes these activities as CE for RNs. This recognition does not imply that AORN or the American Nurses Credentialing Center approves or endorses products mentioned in the activity. http://dx.doi.org/10.1016/j.aorn.2015.02.010 ª AORN, Inc, 2015
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CLINICAL ISSUES
2.6
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THIS MONTH Update on perioperative Ebola precautions Key words: Ebola, PPE, airborne, droplet, respirator. Contaminated rigid sterilization containers Key words: rigid sterilization containers, filters, valves, latches, gaskets. Requirements for storing autologous tissue Key words: tissue bank, autologous tissue, US Food and Drug Administration, human cells, delayed replantation or autotransplantation. Storing autologous human vein autografts Key words: autografts, storage media, Tiprotec, X-Vivo 10, American Association of Tissue Banks. Dropped autografts Key words: autograft, pulse lavage, wound classification, dropped grafts, cranial bone graft. Antimicrobial and fluid barrier fabrics Key words: antimicrobial fabrics, fluid barrier fabrics, surgical site infections, scrub attire, surgical attire.
Update on perioperative Ebola precautions QUESTION:
ANSWER:
Have there been any changes to personal protective equipment (PPE) recommendations when caring for a patient with known or suspected Ebola?
Perioperative team members should continue to take standard, contact, droplet, and airborne precautions when caring for a patient with known or suspected Ebola in the OR. The
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Table 1. PPE for Facilities to Follow When Caring for a Patient With Ebola
1,2
Nonscrubbed team members (eg, RN circulator, anesthesia professional) Follow PPE for surgical N95 respiratora Gown: fluid-resistant, extends to midcalf or Coveralls: fluid-resistant, without integrated hood Boot covers: fluid-resistant, extend to midcalf Gloves: double; outer pair has extended cuff Apron: fluid-resistant, extends to midcalf Full face shield and surgical hood No exposed skin!
Use of PAPRs based on risk assessment: Follow PPE for NIOSH-certified PAPRs Gown: fluid-resistant, extends to midcalf or Coveralls: fluid-resistant, without integrated hood Boot covers: fluid-resistant, extend to midcalf Gloves: double; outer pair has extended cuff Apron: fluid-resistant, extends to midcalf PAPR integrated with full face shield and hood No exposed skin! Scrubbed team members
a
Follow PPE for surgical N95 respirator Sterile surgical gown for AAMI’s level 4 criteria3 Boot covers: fluid-resistant, extend to midcalf Gloves: double, sterile Apron: fluid-resistant, extends to midcalf under sterile gown Full face shield and surgical hood or Surgical helmet system Full face shield and surgical hood or Surgical helmet system No exposed skin!
Do not follow PPE for PAPRs PAPRs should not be used where there is a sterile field during an operative or other invasive procedure.4 Follow PPE for surgical N95 respirator.
Sterile processing team members Option 1: Follow PPE for surgical N95 respiratora Gown5: fluid-resistant, extends to midcalf or Coveralls: fluid-resistant, without integrated hood Boot covers5: fluid-resistant, extend to midcalf Gloves: double; outer pair are general-purpose utility gloves with a cuff that extends beyond the cuff of the gown5 Apron: fluid-resistant, extends to midcalf Full face shield and surgical hood No exposed skin!
Option 2: Follow PPE for NIOSH-certified PAPRs Gown5: fluid-resistant, extends to midcalf or Coveralls: fluid-resistant, without integrated hood Boot covers5: fluid-resistant, extend to midcalf Gloves: double; outer pair are general-purpose utility gloves with a cuff that extends beyond the cuff of the gown5 Apron: fluid-resistant, extends to midcalf PAPR integrated with full face shield and hood No exposed skin!
PPE ¼ personal protective equipment; PAPRs ¼ powered air purifying respirators; NIOSH ¼ National Institute for Occupational Safety and Health; AAMI ¼ Association for the Advancement of Medical Instrumentation. a NIOSH-approved N95 respirator that has also been cleared by the US Food and Drug Administration. References 1. Infection prevention and control recommendations for hospitalized patients with known or suspected Ebola virus disease in U.S. hospitals. Centers for Disease Control and Prevention. http://www.cdc.gov/vhf/ebola/hcp/infection-prevention-and-control-recommendations.html. Updated January 9, 2015. Accessed January 16, 2015. 2. Guidance on personal protective equipment to be used by healthcare workers during management of patients with Ebola virus disease in U.S. hospitals, including procedures for putting on (donning) and removing (doffing). Centers for Disease Control and Prevention. http://www.cdc .gov/vhf/ebola/hcp/procedures-for-ppe.html. Updated October 20, 2014. Accessed January 16, 2015. 3. Guideline for sterile technique. In: Guidelines for Perioperative Practice. Denver, CO: AORN, Inc; 2015:67-96. 4. NPPTL Respirator Trusted-Source Information Section 1: NIOSH-Approved Respirators e What are they? How can they be identified? Where can I get them? http://www.cdc.gov/niosh/npptl/topics/respirators/ disp_part/respsource1.html. Accessed January 16, 2015. 5. Guideline for cleaning and care of surgical instruments and powered equipment. In: Guidelines for Perioperative Practice. Denver, CO: AORN, Inc; 2015:615-650.
Centers for Disease Control and Prevention (CDC) issued updated personal protective equipment (PPE) requirements in the Infection Prevention and Control Recommendations for 576 j AORN Journal
Hospitalized Patients With Known or Suspected Ebola Virus Disease in U.S. Hospitals.1 According to the CDC, when following facility requirements for PPE, perioperative
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Clinical Issues
personnel should have no exposed skin.2 See Table 1 for PPE recommendations in the perioperative setting.
http://www.cdc.gov/vhf/ebola/hcp/procedures-for-ppe.html. Updated October 20, 2014. Accessed January 16, 2015.
References
Amber Wood, MSN, RN, CNOR, CIC
1. Infection prevention and control recommendations for hospitalized patients with known or suspected Ebola virus disease in U.S. hospitals. Centers for Disease Control and Prevention. http://www.cdc .gov/vhf/ebola/hcp/infection-prevention-and-control-recommendations .html. Updated January 9, 2015. Accessed January 16, 2015. 2. Guidance on personal protective equipment to be used by healthcare workers during management of patients with Ebola virus disease in U.S. hospitals, including procedures for putting on (donning) and removing (doffing). Centers for Disease Control and Prevention.
Perioperative Nursing Specialist AORN Center for Nursing Practice
Contaminated rigid sterilization containers QUESTION: I heard that a European study showed bacterial growth on rigid sterilization containers after sterilization. If so, does this present a risk of infection for the patient?
ANSWER: Dunkelberg and Fleitmann-Glende1 examined the microbial growth and barrier properties of 216 sterilization containers after sterilization in a laboratory environment. The containers were obtained from central sterile supply departments from four different hospitals in Germany. The researchers theorized that the integrity of sterilization containers could be affected by repeated use and sterilization; the seal of the container could be weakened by exposure to heat and mechanical stress, causing rivets, bolts, and nuts to loosen or the latching mechanism that secures the lid to the base to deteriorate; and a risk of contamination from an exchange of air between the lower pressure inside of the container and the higher pressure in the environment could result, as might occur when the container is transferred to an area with positive pressure such as the OR. The containers used in the study included containers with paper filters, reusable textile filters, and permanent plastic filters. The researchers placed uncovered thermoresistant plates filled with Sabouraud agar in the base of the containers before sterilizing them. To determine the microbial barrier function of the containers, they exposed the containers to an aerosol suspension of Saccharomyces cerevisiae after sterilization. To determine the potential for microbial contamination caused by changes in atmospheric pressure, the researchers exposed 12 containers to 24 different pressure changes followed by a microbial aerosol of S cerevisiae.
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The researchers found that only two of 11 containers with paper filters and nine of 79 containers with textile filters lacked microbial growth; however, 14 of 15 containers with permanent plastic filters showed no growth. Notably, the contaminated containers showed visible defects, including fissured or loosened lid seals. The researchers found that only four of the 12 containers exposed to the pressure changes maintained a high microbial barrier efficacy. The researchers demonstrated the need for challenging the effectiveness of the microbial barrier of rigid sterilization containers and a method to do so. Although the generalizability of this study may be limited because the containers used by the researchers may differ from those used in the United States, the results of the study emphasize the importance of a quality assurance program that includes careful and periodic inspections of the containers before use in a manner according to the manufacturer’s written instructions for use.2 Personnel should inspect containers to ensure the valves work freely and the sealing surfaces and edges are aligned correctly and are free from dents or chips3; check gaskets to ensure they are pliable, securely fastened, and without breaks or cuts3; inspect filter retention mechanisms and fasteners (eg, screws, rivets) to verify they are secure and functioning correctly; check filter materials for integrity and freedom from holes3; and repair or replace defective containers. The researchers emphasized that although there was no evidence that failure of a sterilization container had caused patient infection,
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the results of their study strongly support the need for monitoring the integrity of packaging systems at least annually.
Sharon A. Van Wicklin, MSN, RN, CNOR, CRNFA, CPSN-R, PLNC
References 1. Dunkelberg H, Fleitmann-Glende F. Measurement of the microbial barrier effectiveness of sterilization containers in terms of the log reduction value for prevention of nosocomial infections. Am J Infect Control. 2006;34(5):285-289. 2. Guideline for sterilization. In: Guidelines for Perioperative Practice. Denver, CO: AORN, Inc; 2015:665-692. 3. Association for the Advancement of Medical Instrumentation (AAMI). ANSI/AAMI ST79:2010/ A4:2013: Comprehensive Guide to Steam Sterilization and Sterility Assurance in Health Care Facilities. Arlington, VA: AAMI; 2013.
Perioperative Nursing Specialist AORN Nursing Department
Requirements for storing autologous tissue QUESTION: Is it true that if a facility stores autologous tissue, the facility must be registered with the US Food and Drug Administration (FDA) as a tissue bank?
ANSWER: A facility that handles autologous tissue for delayed replantation or autotransplantation within the same facility is not required to register with the US Food and Drug Administration (FDA) as a tissue establishment (ie, tissue bank) and is not required to follow the requirements of the FDA regulations for cellular and tissue-based products (x21 CFR Part 1271).1,2 Facilities or health care organizations that handle autologous tissue are required to recover, process, package, label, store, track, and replant or autotransplant the tissue in a manner that minimizes microbial growth, prevents replantation or autotransplantation to the wrong patient, and reduces the risk for errors.1 Although the regulation defines manufacturing to include recovery, processing, storage, labeling, packaging, or distribution of any human cell or tissue,1,3 the FDA considers most procedures related to autologous tissue to be a single procedure encompassed within the element of storage. The reader can refer to section 1271.15(b), of the regulation, in which storage of autologous tissue is exempt if replantation or autotransplantation will occur in the facility where the recovery took place.2 Similarly, packaging and labeling of the autologous tissue can be encompassed within the exception for storage.2 Freezing autologous tissue as a method of storage does not, in
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itself, require the facility to register with the FDA as a tissue establishment.2 In addition, the FDA has interpreted the “same surgical procedure” language in its final rule to include recovery and storage before replantation or autotransplantation.1 Retaining autologous tissue to be used in a subsequent application for the same patient is exempt from registration because the two applications are essentially a single, continuous procedure.2 The facility is required to register with the FDA if autologous tissue handling includes steps to process the autograft that require specific manufacturing controls to decontaminate the tissue (eg, subjecting the autograft to a steam sterilization process).2 If autologous tissueehandling functions are expanded to include distribution of the autograft to another facility located at a different address, registration and listing with the FDA using Form FDA 3356 is required.1
References
1. x21 CFR 1271: Human cells, tissues, and cellular and tissue-based products. April; 2013. US Food and Drug Administration. http:// www.accessdata.fda.gov/scripts/cdrh/cfdocs/cfcfr/CFRSearch .cfm?CFRPart¼1271. Accessed December 6, 2014. 2. Human Cells, Tissues, and Cellular and Tissue-Based Products; Establishment Registration and Listing. Fed Regist. 2001;66(13): 5447-5469. http://www.gpo.gov/fdsys/pkg/FR-2001-01-19/pdf/ 01-1126.pdf. Accessed December 6, 2014. 3. Current Good Tissue Practice (CGTP) and Additional Requirements for Manufacturers of Human Cells, Tissues, and Cellular and TissueBased Products (HCT/ Ps). Silver Spring, MD: US Food and Drug Administration; 2012.
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May 2015, Volume 101, No. 5
Clinical Issues
Sharon A. Van Wicklin, MSN, RN, CNOR, CRNFA, CPSN-R, PLNC
Scott A. Brubaker, CTBS Chief Policy Officer American Association of Tissue Banks McLean, VA
Perioperative Nursing Specialist AORN Nursing Department
Storing autologous human vein autografts QUESTION: How should autologous human vein autografts be stored?
ANSWER: The American Association of Tissue Banks recommends that autologous vein autografts be stored between 32 F to 50 F (0 C to 10 C) for no longer than 14 days.1 Refrigerated veins should be submerged in storage medium, and the storage medium should be changed every 72 hours using sterile technique.2 Keeping the veins submerged helps prevent dehydration of the tissue.2 Changing the storage medium every 72 hours may help prevent microbial growth and improve vein autograft viability.3,4 Three studies provide moderate evidence that regularly changing the storage medium of human vein autografts may extend viability.5-7 To investigate the effect of storing saphenous vein segments in normal saline solution compared with TiProtec solution, Wilbring et al5 isolated 19 saphenous vein segments from patients undergoing coronary artery bypass grafting with autologous saphenous veins in a university heart center between October 2008 and March 2010. The veins were extracted without the use of electrocoagulation. Each vein segment was divided into two parts and placed in either normal saline solution or TiProtec. The vein segments were stored at 39.2 F (4 C) for 96 hours and then examined. The researchers found that the vessel function was significantly reduced after 24 hours of cold storage in normal saline solution. After 96 hours of storage in normal saline solution, there was minimal to no vessel function remaining. Vascular function of the vein segments stored in TiProtec was significantly better preserved. The researchers concluded that cold storage of venous grafts was feasible for as long as 96 hours when the grafts were
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stored in TiProtec solution; however, storage of venous grafts in normal saline solution was not recommended beyond 24 hours.5 Molnar et al6 conducted a quasi-experimental study to investigate the passive and active biomechanical properties of 72 saphenous vein segments remaining after coronary bypass grafting from 32 patients in a university medical center. The vein segments were divided into eight testing groups: fresh, examined immediately after recovery; stored in normal Krebs-Ringer solution at 32 F to 39.2 F (0 C to 4 C), examined one week after recovery; stored in normal Krebs-Ringer solution at 32 F to 39.2 F (0 C to 4 C), examined two weeks after recovery; stored in X-Vivo 10TM at 32 F to 39.2 F (0 C to 4 C), examined one week after recovery; stored in X-Vivo 10 at 32 F to 39.2 F (0 C to 4 C), examined two weeks after recovery; stored in X-Vivo 10 at 32 F to 39.2 F (0 C to 4 C), examined three weeks after recovery; stored in X-Vivo 10 at 32 F to 39.2 F (0 C to 4 C), examined four weeks after recovery; or cryopreserved at -220 F (-140 C), examined three weeks after recovery. Researchers thawed the cryopreserved samples by immersion in warm (98.6 F [37 C]) Krebs-Ringer solution. Biomechanical testing showed that the vein segments stored in Krebs-Ringer solution lost their ability to dilate and contract within one week. The segments stored in X-Vivo 10 preserved their contractility after one week, and contractility slowly decreased during the fourweek study period. There was a slight decrease in vessel wall thickness, but the lumen diameter was not affected. The elastic parameters were almost identical to the fresh segment; however, AORN Journal j 579
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the cryopreserved segments narrowed, the vessel wall thickened, and contractility diminished. The researchers concluded that storage in X-Vivo 10 helped to preserve the passive and active biomechanical properties of human saphenous vein segments. They also concluded that maintaining these properties could be expected to improve stored autologous vein graft viability.6 Baumann et al7 used light transmission and scanning electron microscopy to investigate the effects of various methods of vein preparation on endothelial and smooth muscle cells in cephalic veins from dogs. The researchers removed the veins, divided them into 10-cm segments (N ¼ 10), placed them in one of three solutions (ie, autologous blood [n ¼ 3], Plasma-Lyte [n ¼ 2], or Plasma-Lyte with 0.6 mg/mL papaverine [n ¼ 5]) containing heparin 10,000 units/L, and then stored them at 50 F (10 C) for either five minutes or one hour. The researchers found the vein wall was extremely sensitive to dissection, manipulation, or introduction of fixative solutions and reacted to such stimulations with severe contraction that not only diminished the luminal diameter but also resulted in the protrusion of endothelial cells into the lumen and the formation of cytoplasmic extensions of medial smooth muscle cells. The veins stored in autologous blood demonstrated the greatest amount of vessel wall contraction and endothelial cell loss. Veins stored in Plasma-Lyte for five minutes showed few contractions; however, after one hour, there were contractions and some endothelial cell loss. Veins stored in Plasma-Lyte supplemented with papaverine had the most relaxed appearance and minimal endothelial cell loss. The researchers recommended mitigating damaging vein graft contractions by using gentle surgical dissection to avoid vein spasm; instituting and intermittently repeating (ie, every two to three minutes) gentle perfusion with a solution of Plasma-Lyte supplemented with papaverine immediately after exposure of the distal end of the vein and while the rest of the vein is removed; avoiding the use of autologous blood for vein immersion, storage, or distention; and monitoring the pressure used to check vein grafts for leaks at less than 100 mmHg. The researchers concluded that attention to the details of vein dissection, preparation, and storage medium could lead to a significant improvement in endothelial preservation and subsequent patency rates after cardiac bypass procedures.7 There are gaps in the literature related to best practices for storage of autologous veins, and further research is warranted.
the Lonza Group, Walkersville, MD. Plasma-Lyte is a registered trademark of Baxter Healthcare, Deerfield, IL.
References 1. Standards for Tissue Banking. McLean, VA: American Association of Tissue Banks; 2012. 2. Guideline for autologous tissue management. In: Guidelines for Perioperative Practice. Denver, CO: AORN, Inc; 2015:187-238. 3. Robb EC, Bechmann N, Plessinger RT, Boyce ST, Warden GD, Kagan RJ. Storage media and temperature maintain normal anatomy of cadaveric human skin for transplantation to full-thickness skin wounds. J Burn Care Rehabil. 2001;22(6):393-396. 4. Cram AE, Domayer MA. Short-term preservation of human autografts. J Trauma. 1983;23(10):872-873. 5. Wilbring M, Tugtekin SM, Zatschler B, et al. Preservation of endothelial vascular function of saphenous vein grafts after long-time storage with a recently developed potassium-chloride and N-acetylhistidine enriched storage solution. Thorac Cardiovasc Surg. 2013;61(8):656-662. 6. Molnar GF, Nemes A, Kekesi V, Monos E, Nadasy GL. Maintained geometry, elasticity and contractility of human saphenous vein segments stored in a complex tissue culture medium. Eur J Vasc Endovasc Surg. 2010;40(1):88-93. 7. Baumann FG, Catinella FP, Cunningham JN Jr, Spencer FC. Vein contraction and smooth muscle cell extensions as causes of endothelial damage during graft preparation. Ann Surg. 1981;194(2):199-211.
Sharon A. Van Wicklin, MSN, RN, CNOR, CRNFA, CPSN-R, PLNC Perioperative Nursing Specialist AORN Nursing Department
Scott A. Brubaker, CTBS Chief Policy Officer American Association of Tissue Banks McLean, VA
Editor’s notes: TiProtec is a registered trademark of Dr Franz K€ohler Chemie, Bensheim, Germany. X-Vivo 10 is a trademark of
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Clinical Issues
Dropped autografts QUESTION: Recently, one of our surgeons dropped the autologous cranial bone flap during the replantation procedure and then replanted the contaminated autograft. How should this event have been managed?
ANSWER: The collective evidence indicates there are instances in which autologous grafts necessary for replantation or autotransplantation have been contaminated or dropped on the OR floor and that the infection rate associated with replantation or autotransplantation of contaminated grafts is low.1-3 Various processes for decontaminating the graft have been implemented; however, there is no consistent protocol for managing the event.1-3 Further research is warranted on best practices for decontaminating contaminated autologous grafts and determining methods to prevent the graft from being dropped or contaminated. If an autologous graft intended for replantation or autotransplantation is dropped or contaminated during the procedure, a multidisciplinary team consisting of the surgeon, perioperative RN, and infection preventionist should conduct a risk assessment to consider the benefits and potential harms associated with replantation or autotransplantation of the contaminated autograft compared with other treatment options (eg, discarding the graft and using artificial material).4 If a decision is made to replant or autotransplant the contaminated autograft, the following steps should be taken: Rinse the contaminated graft in sterile normal saline solution to remove surface debris and contaminants. Rinsing the dropped autograft in normal saline solution may be sufficient to remove surface debris and contaminants.4 In a randomized controlled trial conducted to determine the amount of contamination that occurred when a bone graft was dropped on the floor, Presnal and Kimbrough5 concluded that replanting a dropped bone graft was acceptable without extensive, and potentially damaging, efforts to disinfect the graft.5 Use pulsatile lavage at low-pressure settings (eg, 6 psi to 14 psi)6,7 and use sterile normal saline solution for more thorough cleansing of contaminated bone grafts if indicated (eg, adherent debris is present).4 Pulsatile lavage may be useful for removing soil or microorganisms that are difficult to kill or remove with rinsing, such as heavy contamination and debris embedded in the tissue (eg, from a traumatic accident). There is no agreement in the literature as to the absolute definition of
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low-pressure versus high-pressure lavage.6 Bhandari et al8 found that high-pressure lavage (ie, 70 psi) may damage the bone and carry surface contaminants deeper into the bone. In another study, Bhandari et al6 found that low-pressure lavage (ie, 14 psi) was as effective as high-pressure lavage in removing adherent bacteria from bone and preserved the bony architecture when applied within three hours of contamination. Use a separate sterile field for decontaminating the dropped graft and exercise care to prevent splashing onto the primary sterile field.4 Using a separate sterile field may help prevent contamination of the primary sterile field by microorganisms on the contaminated autograft and from droplets caused by rinsing or irrigation fluid used to decontaminate the graft.4 Implementing corrective actions to maintain the sterility of the surgical field may prevent or reduce microbial contamination and help minimize the patient’s risk of surgical site infection.4 Implement corrective actions as necessary to maintain the sterility of the primary sterile field (eg, changing gowns and gloves after pulsatile lavage of the contaminated graft).4 Preventing or reducing microbial contamination helps minimize the patient’s risk of surgical site infection.4 Change the wound classification to Class III, Contaminated.4,9 Replantation or autotransplantation of a contaminated autograft constitutes a major break in sterile technique. According to the Centers for Disease Control and Prevention surgical wound classification system, a surgical wound with a major break in sterile technique is classified as Class III, Contaminated.9 Document the event in a variance report.4 Variance reports document the steps taken during the event and provide a mechanism for alerting infection preventionists to the need for surveillance.4 Variance reports also can be useful for quality improvement activities.4 Conduct a debriefing session and a root cause analysis with members of the surgical team and other individuals who may be helpful in providing a critical analysis and determining the factors that contributed to the event and methods to prevent its recurrence.4 Implementing a process of debriefing and root cause analysis after the event may help prevent future incidents by examining underlying factors and system flaws that may have contributed to the event and may respond to analysis and correction.10
In addition, the following actions may be taken: Add antibiotics or antiseptics to the solution used to rinse the autograft.4 Adding antibiotics or antiseptics to the irrigation solution may be unnecessary or potentially harmful. Antibiotics
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and antiseptics may not have been validated for use in irrigation solutions and may therefore pose a risk to the patient. Tissue proteins may inhibit the effectiveness of the antibiotics.11 Some antibiotics, such as bacitracin,12 have not been found to be effective for eliminating bone contaminants, whereas others, such as rifamycin,13 have been found to be effective for decontaminating contaminated bone grafts. Antiseptic solutions are intended for external use and may be ineffective or toxic when used internally. A povidone-iodine solution has been found to provide effective bone decontamination14; however, the antibacterial activity of 10% povidone-iodine solution is directly related to the level of contamination and the duration of exposure to the contaminated bone.15 Lacey16 found that the antibacterial effect of a povidone-iodine solution was inactivated in the presence of blood. Kaysinger et al17 found that 0.5%, 5%, and 50% povidone-iodine solutions and 1.5% and 3% hydrogen peroxide (H2O2) solutions were cytotoxic to osteoblasts. Bhandari et al6 found that a povidone-iodine solution and bacitracin decreased the number of osteoclasts and impaired osteoblast function. When used to disinfect explanted bone, Sch€ ultke et al18 found that H2O2 was ineffective. Studies have shown chlorhexidine gluconate to be both effective14 and ineffective13 for decontaminating contaminated bone grafts and it has also been found to be toxic to bone cells, even at very low concentrations (ie, 1%).12 Using chemical disinfectants (eg, glutaraldehyde) on bone may cause tissue damage or the disinfectants may be difficult to remove.4 Take samples for culture before and after decontamination of the graft to determine the identity of the contaminating microorganism and the level of contamination.4 Tissue cultures may be helpful in determining whether the dropped autograft was contaminated before or after decontamination measures were implemented.4 The results of the cultures may be useful in guiding treatment after replantation.4 Consult with an infection preventionist to assess the benefits versus harms of implementing postoperative broad-spectrum antibiotic prophylaxis therapy.4 Send the autograft to a tissue bank for decontamination and processing.4 Some tissue banks provide services to decontaminate autologous bone skull flaps when contamination is suspected (ie, dropped autograft, head trauma, positive culture result).4 The contaminated graft should not be subjected to the steam sterilization process,4 which denatures bone proteins, may severely damage the bone structure, and increases the potential for bone resorption and infection.18 The steam sterilization process has not been validated for use with human tissue. A significantly high rate of postoperative graft infections has been found when cranial bone grafts subjected to the steam sterilization process were used for cranial reconstruction.19
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References
1. Jankowitz BT, Kondziolka DS. When the bone flap hits the floor. Neurosurgery. 2006;59(3):585-589. 2. Kang L, Mermel LA, Trafton PG. What happens when autogenous bone drops out of the sterile field during orthopaedic trauma surgery. J Orthop Trauma. 2008;22(6):430-431. 3. Centeno RF, Desai AR, Watson ME. Management of contaminated autologous grafts in plastic surgery. Eplasty. April 22, 2008;8:e23. 4. Guideline for autologous tissue management. In: Guidelines for Perioperative Practice. Denver, CO: AORN, Inc; 2015:187-238. 5. Presnal BP, Kimbrough EE. What to do about a dropped bone graft. Clin Orthop Relat Res. 1993;296:310-311. 6. Bhandari M, Schemitsch EH, Adili A, Lachowski RJ, Shaughnessy SG. High and low pressure pulsatile lavage of contaminated tibial fractures: an in vitro study of bacterial adherence and bone damage. J Orthop Trauma. 1999;13(8):526-533. 7. Hirn M, Laitinen M, Pirkkalainen S, Vuento R. Cefuroxime, rifampicin and pulse lavage in decontamination of allograft bone. J Hosp Infect. 2004;56(3):198-201. 8. Bhandari M, Adili A, Lachowski RJ. High pressure pulsatile lavage of contaminated human tibiae: an in vitro study. J Orthop Trauma. 1998;12(7):479-484. 9. Mangram AJ, Horan TC, Pearson ML, Silver LC, Jarvis WR. Guideline for Prevention of Surgical Site Infection, 1999. Centers for Disease Control and Prevention (CDC) Hospital Infection Control Practices Advisory Committee. Am J Infect Control. 1999;27(2): 97-132; quiz 133-134; discussion 96. 10. Makary MA, Holzmueller CG, Sexton JB, et al. Operating room debriefings. Jt Comm J Qual Patient Saf. 2006;32(7):407-410. 357. 11. Sample Procedure: Handling Autologous Bone Skull Flaps. Version 14. McLean, VA: American Association of Tissue Banks; 2012. 12. Bhandari M, Adili A, Schemitsch EH. The efficacy of low-pressure lavage with different irrigating solutions to remove adherent bacteria from bone. J Bone Joint Surg Am. 2001;83-A(3):412-419. 13. Yaman F, Unl€u G, Atilgan S, Celik Y, Ozekinci T, Yaldiz M. Microbiologic and histologic assessment of intentional bacterial contamination of bone grafts. J Oral Maxillofac Surg. 2007;65(8): 1490-1494. 14. Bruce B, Sheibani-Rad S, Appleyard D, et al. Are dropped osteoarticular bone fragments safely reimplantable in vivo? J Bone Joint Surg Am. 2011;93(5):430-438. 15. Soyer J, Rouil M, Castel O. The effect of 10% povidone-iodine solution on contaminated bone allografts. J Hosp Infect. 2002; 50(3):183-187. 16. Lacey RW. Antibacterial activity of povidone iodine towards nonsporing bacteria. J Appl Bacteriol. 1979;46(3):443-449. 17. Kaysinger KK, Nicholson NC, Ramp WK, Kellam JF. Toxic effects of wound irrigation solutions on cultured tibiae and osteoblasts. J Orthop Trauma. 1995;9(4):303-311. 18. Sch€ultke E, Hampl JA, Jatzwauk L, Krex D, Schackert G. An easy and safe method to store and disinfect explanted skull bone. Acta Neurochir (Wien). 1999;141(5):525-528. 19. Matsuno A, Tanaka H, Iwamuro H, et al. Analyses of the factors influencing bone graft infection after delayed cranioplasty. Acta Neurochir (Wien). 2006;148(5):535-540.
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May 2015, Volume 101, No. 5
Clinical Issues
Sharon A. Van Wicklin, MSN, RN, CNOR, CRNFA, CPSN-R, PLNC
Scott A. Brubaker, CTBS Chief Policy Officer American Association of Tissue Banks McLean, VA
Perioperative Nursing Specialist AORN Nursing Department
Antimicrobial and fluid barrier fabrics QUESTION: Are fabrics available that can prevent bacteria from being retained on the surface of the material? If so, can using these fabrics for scrub attire provide protection for perioperative personnel and also help prevent surgical site infections?
ANSWER: According to the AORN “Guideline for surgical attire,”1 “Scrub attire may be made of antimicrobial fabric.”1(p101) There is emerging evidence regarding the use of fabrics with fluid barrier and antimicrobial properties to prevent bacteria and fungi from adhering to the fabric.1 Bearman et al2 conducted a randomized controlled trial to determine the effectiveness of an antimicrobial, fluid barrier scrub fabric for reducing the bacterial burden on the hands of and scrub attire worn by health care workers in an intensive care unit setting of an academic medical center. All study participants (N ¼ 30) were randomly assigned to wear either traditional scrub attire or scrub attire made of antimicrobial, fluid barrier fabric during a clinical shift during a four-week period. The researchers took cultures from the abdominal and leg pockets of the scrub attire at the beginning and end of the shifts. Notably, the researchers did not specify the length of the clinical shifts. The researchers found a significant difference in the number of methicillin-resistant Staphylococcus aureus colony-forming units in both the leg and abdominal area of the antimicrobial, fluid barrier scrub attire compared with traditional scrub attire at the beginning and end of shifts. The researchers concluded that when combined with infection prevention strategies, such as hand hygiene and the use of personal protective equipment,
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the use of antimicrobial, fluid barrier scrub attire may limit the bacterial burden of the inanimate environment.2 In a quasi-experimental study to examine whether antibacterial finishes on fabric could effectively reduce the presence of bacteria on fabric used for health care workers’ uniforms, Chen-Yu et al3 found that the antibacterial finishes provided a significant reduction of S aureus and Klebsiella pneumoniae. The researchers concluded that adding antibacterial finishes to fabric was an effective method of reducing bacterial contamination. Mariscal et al4 conducted a quasi-experimental study to evaluate the action of a commercially available antimicrobial textile on 33 strains of bacteria. They found that the antimicrobial fabric significantly reduced the numbers of four reference microorganisms (ie, Escherichia coli, Pseudomonas aeruginosa, Morganella morgannii, S aureus) compared with the numbers of reference microorganisms on the control fabric. Sun et al5 quantitatively and qualitatively evaluated fabrics treated with antimicrobial chemicals in a nonexperimental study. The researchers found that even after 50 washings, the treated fabrics exhibited antibacterial properties against gram-negative (eg, Escherichia coli) and gram-positive (eg, S aureus) bacteria and fungi (eg, Candida albicans) within a two-minute contact time. Despite the biocidal properties of the antimicrobial fabric, the researchers found it was nontoxic to human skin. They concluded that antimicrobial fabrics were effective and suitable for medical use. In a randomized crossover trial conducted to determine whether applying an antimicrobial coating to scrub attire worn by nurses and patient care technicians working in the adult intensive and intermediate care units of a university medical center would reduce the overall rate of scrub contamination with pathogenic
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bacteria (eg, S aureus, Enterococcus species, gram-negative bacteria), Boutin et al6 cultured the front of 720 scrub tops worn by 90 health care workers during the last four hours of a 12-hour shift. The researchers found that 30% (n ¼ 217) of the scrub tops were contaminated with pathogenic bacteria. The treated scrub attire showed a contamination rate of 30% (n ¼ 108), and the untreated scrub attire showed a contamination rate of 30.3% (n ¼ 109). The researchers concluded the antimicrobial coating did not prevent bacterial contamination of scrub attire. In a quasi-experimental study to evaluate the bactericidal effects of woven and nonwoven fabrics coated with a hydroxyapatite-binding silver/titanium dioxide ceramic composite, Kasuga et al7 found that bacterial cell counts of S aureus and E coli on both the woven and nonwoven fabrics decreased to less than 2-log10 colony-forming units/mL within six hours and were undetectable at the end of the 18-hour incubation period. Bacterial cell counts of P aeruginosa could not be detected after three to six hours. In a nonexperimental study of silver-impregnated scrub attire, Gross et al8 compared the contamination rates of traditional ambulance service clothing and silver-impregnated ambulance service clothing worn by 10 emergency workers. The samples were taken from the right sleeves and front of the jackets and the right thighs of the pants before use and at the end of the third and seventh working days. Notably, the researchers did not specify the length of the working day. The researchers found no significant difference in the amount of contamination between the two clothing types; however, they recommended verifying the result of the study with a larger sample size.
5. Sun G, Qian L, Xu X. Antimicrobial and medical-use textiles. Textile Asia. 2001;32(9):33-35. 6. Boutin MA, Thom KA, Zhan M, Johnson JK. A randomized crossover trial to decrease bacterial contamination on hospital scrubs. Infect Control Hosp Epidemiol. 2014;35(11):1411-1413. 7. Kasuga E, Kawakami Y, Matsumoto T, et al. Bactericidal activities of woven cotton and nonwoven polypropylene fabrics coated with hydroxyapatite-binding silver/ titanium dioxide ceramic nanocomposite “Earth-plus.” Int J Nanomed. 2011;6:1937-1943. 8. Gross R, H€ubner N, Assadian O, Jibson B, Kramer A. Pilot study on the microbial contamination of conventional vs. silver-impregnated uniforms work by ambulance personnel during one week of emergency medical service. GMS Krankenhhyg Interdiszip. 2010; 5(2):1-16. 9. Bauer J, Kowal K, Tofail SAM, Podbielska H. MRSA-resistant textiles. In: Tofail SAM, ed. Biological Interactions with Surface Charge in Biomaterials. Cambridge, England: RSC Publishing; 2012:193-207. 10. Rajendran R, Radhai R, Kotresh TM, Csiszar E. Development of antimicrobial cotton fabrics using herb loaded nanoparticles. Carbohydr Polym. 2013;91(2):613-617.
Sharon A. Van Wicklin, MSN, RN, CNOR, CRNFA, CPSN-R, PLNC Perioperative Nursing Specialist AORN Nursing Department
Incorporating textile technologies into the material used for scrub attire and other garments worn by health care personnel may help prevent microorganisms from being acquired and retained on the fabric and reduce the risk of transmission of microorganisms that may cause surgical site infections.2-10 This issue warrants further research.
References 1. Guideline for surgical attire. In: Guidelines for Perioperative Practice. Denver, CO: AORN, Inc; 2015:97-120. 2. Bearman GM, Rosato A, Elam K, et al. A crossover trial of antimicrobial scrubs to reduce methicillin-resistant Staphylococcus aureus burden on healthcare worker apparel. Infect Control Hosp Epidemiol. 2012;33(3):268-275. 3. Chen-Yu JH, Eberhardt DM, Kincade DH. Antibacterial and laundering properties of AMS and PHMB as finishing agents on fabric for health care workers’ uniforms. Clothing Text Res J. 2007;25(3):258-272. 4. Mariscal A, Lopez-Gigosos RM, Carnero-Varo M, FernandezCrehuet J. Antimicrobial effect of medical textiles containing bioactive fibres. Eur J Clin Microbiol Infect Dis. 2011;30(2):227-232.
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Lisa Spruce, DNP, RN, CNS-CP, ACNS, ACNP, ANP, CNOR Director, Evidence-Based Perioperative Practice AORN Nursing Department
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LEARNER EVALUATION
Continuing Education: Clinical Issues 2.6
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T
his evaluation is used to determine the extent to which this continuing education program met your learning needs. The evaluation is printed here for your convenience. To receive continuing education credit, you must complete the online Learner Evaluation at http://www.aorn.org/CE. Rate the items as described below.
PURPOSE/GOAL To provide the learner with knowledge of standard Ebola precautions related to personal protective equipment as well as AORN’s guidelines related to contaminated rigid sterilization containers, requirements for storing autologous tissue, storing autologous human vein autografts, dropped autografts, and antimicrobial and fluid barrier fabrics.
6.
Will you be able to use the information from this article in your work setting? 1. Yes 2. No
7.
Will you change your practice as a result of reading this article? (If yes, answer question #7A. If no, answer question #7B.)
7A.
How will you change your practice? (Select all that apply) 1. I will provide education to my team regarding why change is needed. 2. I will work with management to change/implement a policy and procedure. 3. I will plan an informational meeting with physicians to seek their input and acceptance of the need for change. 4. I will implement change and evaluate the effect of the change at regular intervals until the change is incorporated as best practice. 5. Other: __________________________________
7B.
If you will not change your practice as a result of reading this article, why? (Select all that apply) 1. The content of the article is not relevant to my practice. 2. I do not have enough time to teach others about the purpose of the needed change. 3. I do not have management support to make a change. 4. Other: __________________________________
8.
Our accrediting body requires that we verify the time you needed to complete the 2.6 continuing education contact hour (156-minute) program: _________________________________
OBJECTIVES To what extent were the following objectives of this continuing education program achieved? 1. Discuss practices that could jeopardize safety in the perioperative area. Low 1. 2. 3. 4. 5. High 2.
Discuss common areas of concern that relate to perioperative best practices. Low 1. 2. 3. 4. 5. High
3.
Describe implementation of evidence-based practice in relation to perioperative nursing care. Low 1. 2. 3. 4. 5. High
CONTENT 4.
5.
To what extent did this article increase your knowledge of the subject matter? Low 1. 2. 3. 4. 5. High To what extent were your individual objectives met? Low 1. 2. 3. 4. 5. High
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