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Let’s begin this article by saying, “You’re right!” Whatever oxygen flow rate you routinely use when anesthetizing your patients, whether it’s determined by careful calculation, by habit, or by setting it where you’re told to set it, “You’re right!” This is one of those wonderful times when there is no universal agreement as to the proper flow rate for the various anesthesia breathing systems. So, the pressure’s off. You’re doing it right. [exhale]
Now that we’ve established there are different right ways to do it, we’re free to consider some new right ways to try. But most of us are reluctant to tamper with the way we run anesthesia because we know anesthesia is part science and part magic. And although we trust the science part, magic can be unpredictable. Before we attempt new right ways to do things, let’s take a look at some of the science behind how to decide the oxygen flow rate for small animal anesthesia.
How much oxygen does the patient need?
The oxygen flowing through an anesthetic gas machine is used to provide the patient with sufficient oxygen and to provide a carrier for the anesthetic agent. Since the oxygen flow meter regulates the flow of oxygen per minute, we need to know how much oxygen our patient uses in a minute. This is called the metabolic oxygen consumption rate per minute. A safe estimate of oxygen consumption for small animal patients is 4 – 7ml/kg/min or 2 – 3ml/lb/min. That means that my 14 pound cat Ellie is consuming around 40ml of oxygen each minute she monitors the dog’s behavior from her perch on the couch.
The kind of breathing circuit makes a difference
In general terms, there are two kinds of anesthesia breathing circuits: rebreathing and non-rebreathing. Rebreathing circuits are most often circle systems that include a carbon dioxide absorber like soda lime. The granular soda lime extracts exhaled carbon dioxide from the circuit which allows the gas to be breathed again or rebreathed. Rebreathing circuits use lower gas flows, which decreases the cost and decreases pollution while it retains moisture and heat for the patient. However the soda lime and unidirectional flow valves of a circle system are significant sources of resistance to breathing. Smaller patients can’t overcome the resistance of a circle system and so you must use a non-rebreathing circuit. Non-rebreathing circuits depend on high oxygen flow to remove exhaled carbon dioxide from the circuit between breaths. High oxygen flow rates are inefficient, expensive, and carry heat and moisture away from the patient. The decision to select a non-rebreathing circuit is often made by the weight of the animal, but it is actually a decision that the patient is too small to overcome the resistance of a rebreathing circuit.
Because of the simplicity of a non-rebreathing circuit, there is little debate over what recommended oxygen flow rate to use. The AAHA recommended flow rate of 200ml/kg/min is generally accepted as appropriate. That flow rate is 33 times more oxygen than is needed to meet a patient’s metabolic oxygen consumption each minute, but that high flow rate assures the patient will not rebreathe any of its exhaled carbon dioxide.
Rebreathing circuits offer a wider selection of flow rates. The same rebreathing circle system can be operated as a closed system, a low-flow system, or a semiclosed system, depending on the selected oxygen flow rate. They do not depend on the position of the pop-off valve. The pop-off valve should always be fully open regardless of the oxygen flow rate.
The closed system flow rate meets the patient’s actual metabolic need for oxygen each minute. It’s a style of anesthesia that demands a thorough understanding of the patient, the anesthetic agent, the gas machine, and the varying degrees of stimulation during the surgical procedure. It’s best suited for anesthesia nerds. Some variation of the semiclosed system is used in most small animal practices. The semiclosed flow rate well exceeds the patient’s metabolic requirement for oxygen, and a significant amount of excess gas is exhausted through the pop-off value. The flow rate traditionally falls within 22 – 44ml/kg/min, most often settling at 30ml/kg/min. Remember that the patient’s metabolic oxygen consumption rate per minute is less than 10ml/kg, so there is little concern that you won’t provide enough oxygen flow.
The vaporizer plays a role in your oxygen flow rate
AAHA also makes recommendations for the oxygen flow rate when using a rebreathing circle system. The AAHA recommendation during the maintenance phase of anesthesia is between 200 and 500ml/min. That flow is not calculated using the weight of the patient; it is the recommended setting of the flow meter. They warn that the gas machine must be leak free for those flows to be effective, and concedes that flows that low may be out of the comfort zone of most of us. However, they may not have considered the tendency for most vaporizers to over deliver at low flow rates. Most vaporizers recommend oxygen flow rates not less than 500ml/min for accurate delivery of anesthetic gas.
Ready to try a new right way?
Whatever oxygen flow rate you routinely use when anesthetizing your patients, whether it’s determined by careful calculation, by habit, or by setting it where you’re told to set it, “You’re right!” Now that we’ve taken a look at some of the science behind how to decide the oxygen flow rate for small animal anesthesia, maybe we’ll try another right way.
For more information, refer to AAHA Anesthesia Guidelines for Dogs and Cats
A special guest blog post by Dr Colin Dunlop.
Colin Dunlop is a Diplomate of the American College of Veterinary Anesthesiologists. His research interests include hypothermia and prevention of anesthesia morbidity and mortality. He consults in anesthesia and critical care for small and large animal practices, biomedical research, and provides Continuing Education programs for veterinarians and veterinary nurses world-wide.
In my spare time I try to run a company that manufactures anesthetic delivery equipment and devices for patient warming, so please understand that some of the information I have included references specific warming devices, some of which we manufacture. The information sheets linked to this post try to fairly assess the capabilities of the various technologies and summarize information we have from published research and data from our in-house testing. Written here is information I wrote in a hypothermia article some years ago. It is simply to highlight the practicality of warming a hypothermic patient using IV fluid. It is written in calories, and a Calorie is the heat required to raise 1 ml water by 1 degree Celsius.
Using the same information, you could also attempt to a very simplistic estimate of heat loss due to humidification of inspired air. Heat conservation mechanisms (the nose!) play a big part, and intubation completely alters them. Recent work shows that the loss of heat of warmed gas from the Y-piece to the distal end of the endotracheal tube is up to 10 degrees Celsius!
Not all IV Fluid Warmers are the same, and some can even be dangerous. Test data from our evaluation of IV Fluid Warmers can be found here: IV Fluid Warmer Comparison. According to our tests, most are ineffective if their performance is tested 200mm downstream from the fluid warmer, which is equivalent to the distance to an anesthetized animal, draped for surgery. Their performance is also affected by IV fluid flow rate – the larger the volume of fluid administered rapidly the less effective they are at warming.
Hanging bags of warm IV fluids in a cold operating room will produce the same kind of result as using an IV Fluid Line Warmer, as illustrated in the evaluation of IV Fluid Warmers linked above.
The one-page guide linked here: Hypothermia Review summarizes information from various sources and includes an idealized graph that show heat loss occurs from the time of premedication, and that substantial heat loss occurs from induction to the time an animal is draped in surgery. Once draped, heat loss tends to stabilize, but warming hypothermic anesthetized animals during surgery is very difficult. In fact even the best forced warm air heating systems, which are the most effective way to deliver large volumes of heat, typically take 45minutes of “contact” before body temp begins to rise. So using forced warm air heating devices to “increase” the body temperature during anesthesia for short procedures is probably not very effective. The rationale for pre-warming and preventing heat loss prior to draping for surgery is where our efforts should be focused.
The white paper linked here: FWAH Review and Cage Warming shows the lag-time for warming and that not all these systems actually raise body temperature. Also this paper makes the case for warming animals after premedication, before induction. Typically, warming animals during recovery, who became hypothermic in surgery animals, takes 1 to 2 hours of technician time. Conversely, keeping animals at 37 degrees Celsius takes less than 45 min of “pre-warming”, and the blankets placed over animals in cages can be re-used.
Finally, use warm fluids effectively in severely hypothermic animals at the end of abdominal or thoracic surgery by pouring large volumes of warm (40 degrees Celsius) into the abdominal or thoracic cavity. Be patient. Wait several minutes before suctioning it out and then repeat this process three or four times, until the body temperature starts to rise. Then close the cavity. At the same time use forced warm air heating, which will further increase the body temperature.
Calculating Calories and Warming With IV Fluids
A calorie (cal) is the amount of heat required to raise 1 ml (or 1 gm) of H2O 1 oC.
The specific heat of animal tissue is 0.83 cal/gm. Therefore a 10 kg dog requires 8,300 cal (8.3 kcal) to raise its temperature 1 oC.
Warming IV fluid administered during surgery:
A 10 kg dog administered IV fluid at 10 ml/kg/hr = 100 ml/hr.
If the fluid is warmed to 44 oC and the dog is 34 oC, then we can deliver:
(44-34oC =) 10 oC x 100 ml/hr = 1000 cal/hr
To Warm the 10 kg at 34 oC dog to 37 oC using IV fluid at 100 ml/hr requires:
(37-34 =) 3oC x 8,300 cal = 25,000 cal (approx) / 1000 cal/hr (from the IV fluid) = 25 hours!
Warming IV fluid may help prevent cold fluid exacerbating heat loss but is not effective for warming severely hypothermic animals.
Respiratory heat loss due to humidification is significant
During inspiration the nose and pharyngeal mucosa transfer heat and moisture to the air which is largely recovered during expiration, thus conserving heat. Air has a low heat capacity (0.24 cal/gm) and a low weight (1.3 gm/l). Saturated air holds 44 mg H2O/L at 37 oC which requires 24 calories. A 10 kg dog taking 20 x 100ml breaths/min ventilates 120 L/hr so requires (24 cal/L x 120 L/hr) = 2880 cal/hr for humidification. Intubation inhibits heat/moisture conservation via the nose, resulting in body temperature loss of about 1/3 oC/hr.
We’re really excited to be exhibiting at the IVECCS / ACVAA conference being held at the Gaylord National Resort and Convention Center in Washington, DC. It’ll be a great time to reconnect with old friends and show off what innovations Advanced Anesthesia Specialists brings to veterinary practice this year. Spoiler Alert – we’re introducing a heated bain circuit this year. Heated! Yeah. We’re excited about it too!
If you’re at the conference, stop by our booth #326 and say “hi”.
dealflow is a quarterly magazine published by the Australian Government Department of Industry and Science. It showcases high-performing, small and medium sized Australian companies supported by the Accelerating Commercialisation element of the Government’s Entrepreneurs’ Programme. The global efforts of Dr Colin Dunlop to improve anesthesia outcomes for small animals are applauded in the current issue of dealflow. The article is reproduced below.
Few companies can claim to help household pets while reducing greenhouse gases, but that’s what Advanced Anesthesia Specialists and its managing director Dr Colin Dunlop are doing.
The company is becoming a global leader in the design, manufacture and service of innovative veterinary anesthesia equipment, and continues to break new frontiers.
Dunlop says the mortality risk of anesthesia for animals under 20 kilograms, such as dogs and cats, is about 500 times higher than for humans. To improve survival rates the company has developed a new integrated anesthesia delivery system with Australian Government commercialization support.
The new system has three components. A Heated Smooth Wall Anesthesia tubing system, which warms the gas delivered to patients. This world-first system was released in Australia in 2014 and in UK and US markets this year. It helps to prevent hypothermia, the commonest complication of anesthesia and surgery.
“Hypothermia occurs in up to 85 per cent of anesthetized human infants and small animals,” Dunlop says.
The second component is the Stingray—the first low-flow, low‑resistance with rapid response rebreathing anesthesia circuit for patients under 20 kilograms. It improves on existing anesthesia technology and recycles exhaled breath, which also helps to reduce the risk of hypothermia.
The Stingray, which will be released to global markets in September this year, also reduces the release of environmentally harmful anesthetic gas into the atmosphere by up to 90 per cent.
The system’s third element is an anesthetic vaporizer which provides early warnings of problems during surgery. Dunlop says this novel system, which is in clinical trial stage, will help fill a gap in anesthetic training among veterinarians and veterinary nurses. It is due for release in mid-2016.
International usability is a vital element of the company’s design work. “We could never afford to design these products just for Australia, as the volume of potential sales here is too small to be cost-effective … we need to design equipment for use around the world,” he says.
Protecting intellectual property is also a company priority. “We have invested a lot of money and time in protecting the IP of our technology and devices and have over 13 families of patents, plus new patent applications lodged,” Dunlop says.
Australian Government Commercialisation Adviser, John Grew, has been assisting the company with its move into markets.
“Colin and his team have addressed the many challenges of refining the design, prototyping and pre-production of their products,” Grew says. “A lot of new IP has been developed and the products now coming to market reflect experience and insight.”
Advanced Anesthesia Specialists is owned by veterinarians and Dunlop’s main focus is to improve the odds for pets undergoing anesthesia in Australia and overseas.
“Our company is making a difference—by developing better, more sophisticated equipment we are improving outcomes for both veterinary staff, their patients and the environment.”
To visit this article in dealflow, click here
To visit website for Advanced Anesthesia Specialists, North America, click here
One of my favorite mental pictures of the cardiovascular system is that of A Pump, Some Pipes, and Fluid. Broken down to this simple picture, it’s easier for me to interpret the information I gather as I monitor anesthetized patients.
The heart is at the top of this minimalist’s view of the cardiovascular system by acting as a pump. Its job is to pump blood around the body. The left side of the heart pumps oxygenated blood from the lungs to the rest of the body. The right side pumps stale blood from the body back to the lungs for a fresh supply of oxygen.
The pipes, of course, are those estimated 60,000 miles of veins and arteries distributed throughout the body. Most blood vessels can alter their size in order to accommodate the necessary flow of blood. When a vessel’s interior grows larger to allow more blood flow, it’s called vasodilation. When it shrinks down to decrease blood flow it’s called vasoconstriction. Under normal circumstances, the vessels automatically vasodilate and vasoconstrict to help regulate blood flow through the body. However, many anesthesia drugs alter the body’s ability to respond automatically in this manner.
It is generally accepted that most domestic animals have blood volumes of about 7% of their body weight (cats have a little lower percentage). That equals about 70ml per kilogram or about 35ml per pound of body weight. That means your 60 pound Labrador has a blood volume of about half a gallon. When you consider that a half gallon of blood is pumped through 60,000 miles of blood vessels, you realize that it can’t be all places all the time. The body is constantly making choices to route blood where it is most needed at any given point in time.
Pressure is the driving force for blood flow through capillaries that supply oxygen to organs and tissues. Blood pressure is needed to propel blood through vascular beds, with priority to those of the brain, heart, lungs and kidneys. When I notice a drop in blood pressure, I immediately run through this simplified picture of the cardiovascular system. Why is the pressure dropping? Is the problem with the pump (ie not pumping hard enough or fast enough)? Is the problem with the pipes (ie vasodilation or positional occlusion of major vessels)? Or is the problem with the fluid (ie blood loss or vascular pooling)? The answers to these questions can help me anticipate a corrective treatment.
Anesthesia guidelines from the American College of Veterinary Anesthesia and Analgesia (ACVAA) and the American Animal Hospital Association (AAHA) urge us to monitor blood pressure during anesthesia, yet specialists say that blood pressure equipment alone is not the main ingredient to a smooth anesthetic event. It’s the anesthetist’s knowledge that provides the greatest margin of safety for the patient.
How many food-related decisions do you make in a day? Ten? Fifty? A hundred?
When Cornell University staff and students were asked this question in a recent study, the average response was fifteen. However, after they answered specific questions about when, what, how much and where they ate, researchers found they actually made an average of 221 food-related decisions each day.
The more decisions we make throughout the day, the harder each choice is for our brains to process. No matter how rational and clear-minded you try to be, you can’t make the estimated 35,000 daily decisions we all make, without paying a biological price. It’s called decision fatigue. Although you are not consciously aware of being tired, each decision you make takes its toll on your mental energy. Eventually our brains look for shortcuts. One shortcut is to become reckless: to act impulsively instead of expending the additional energy to think things through. The other shortcut is to simply do nothing at all.
“Good decision-making is not a trait of the person, in the sense that it’s always there,” says social psychologist Roy F Baumeister. “It’s a state that fluctuates.”
Baumeister, a researcher and professor of psychology at Florida State University says, “The best decision makers are the ones who know when not to trust themselves.”
In high-pressure environments like a veterinary hospital, we are up against two main decision-making difficulties. The first is our own fallible memory and attention, especially when routine matters are overlooked under the strain of more pressing events. In the swirl of sedating a patient, placing a catheter, inducing anesthesia, clipping for surgery, and transporting the patient to the surgery table, it’s easy to overlook the routine task of checking the amount of oxygen left in the tank.
The second difficulty is that we sometimes lull ourselves into deliberately skipping steps, even when we remember them. Even the most competent people have been known to tell themselves that certain steps don’t matter. For example, it’s standard to check a patient’s pulse and heartrate during a physical examination. But being diligent to do both – listen to the heart and feel the pulse – rarely uncovers a worrisome issue. So sometimes we skip one. “It’s never been a problem before,” we tell ourselves. Until one day it is.
Simple checklists can provide protection against these mental failures. According to Atul Gawande, in his book The Checklist Manifesto, checklists remind us of the bare minimum necessary steps in any procedure by making them explicit. They catch mental flaws inherent in all of us – flaws of memory and attention.
A good checklist doesn’t have to be long to be effective. But good checklists are always precise and efficient. They leave zero room for interpretation. Don’t attempt to spell out every single step, instead provide reminders of only the most critical and important steps. Good checklists are, above all, practical.
Once a checklist is written, it’s important to be disciplined and stick with it. No matter how smart, talented or experienced we are chances are we’ll still drop the ball sometimes. And we’re not alone. The fact is, we are all plagued by missed subtleties, overlooked knowledge, and out-right errors. No one is immune to screwing-up. Checklists are absolutely essential for handling the high stakes and complex situations that happen to anesthetists. As Gawande points out in The Checklist Manifesto, simple checklists save time and money. But more importantly, in our line of work, checklists save lives.
Many of us think we’re feeling burn-out with our jobs when we’re actually feeling compassion fatigue. Two terms for the same thing? Here’s how you recognize the difference. Burn-out always arises from dissatisfaction with your work environment. It’s generally because of supervisors, poor working conditions, low pay, and/or the relationships you have with the people at work. Compassion fatigue arises from the work that you do.
Compassion fatigue is a more user friendly term for Secondary Traumatic Stress Disorder, which is nearly identical to Post-Traumatic Stress Disorder (PTSD), except it affects those emotionally affected by the trauma of another. Charles Figley, professor of Disaster Mental Health at Tulane University’s School of Social Work and coauthor of Compassion Fatigue in the Animal Care Community says, “It’s the burden of caring. It’s the psychosocial sadness we take with us. It’s the stress of dispensing compassion.”
The solution to burn-out is pretty straight forward: find another job. However, the residual emotional effects of intense medical experiences such as euthanasia aren’t so easily solved. Dr. Kristin Neff, associate professor in Human Development and Culture at the University of Texas thinks self-compassion is at the heart of relieving compassion fatigue. She says self-compassionate people tend to be gentle with themselves when confronted with painful experiences. When people try to deny or resist their reactions to painful experiences, emotional suffering escalates into stress, frustration and self-criticism.
People who find it easy to be supportive and understanding to others – including their animal patients – often berate themselves for their own self-perceived shortcomings. Research suggests that giving ourselves a break and accepting our imperfections may be the first step toward better health. People who score high on tests of self-compassion have less depression and anxiety, and tend to be happier and more optimistic.
For those low on the self-compassion scale, Dr. Neff suggests a set of exercises — like writing yourself a letter of support, just as you might to a friend you are concerned about. She says to include in the letter a list of your best traits, and add steps you might take to help you feel better about yourself.
“The problem is that it’s hard to unlearn habits of a lifetime,” she says about our tendency to equate self-compassion with self-indulgence. “People have to actively and consciously develop the habit of self-compassion.”
For more information about Dr Kristin Neff’s work in self-compassion, visit http://www.self-compassion.org/
Is there a relationship between exposure to trace concentrations of waste anesthetic gasses and the development of health concerns? After two independent groups analyzed more than seventeen studies, including one well-designed prospective study, the consensus is that there is no risk of adverse health effects to personnel where waste anesthetic gases are scavenged.
Studies published in the late 1960’s and early 1970’s pointed to waste anesthetic gas (WAG) as a direct contributor to everything from fatigue, exhaustion, and headaches to cancer, infertility, spontaneous abortion, and birth defects. These studies resulted in a 1974 National Institute for Occupational Safety and Health (NIOSH) recommendation that waste anesthetic gas be scavenged in all areas. Three years later NIOSH recommended that WAG exposure standards be established.
In the 1980’s, researchers examined the conclusions drawn from these earlier studies. Seventeen studies were examined and all were found to have flaws. They found that the results of these studies could have been influenced by confounding variables such as occupational stress, and exposure to blood, drugs, aerosols or radiation. Even the wording of the questionnaires and the inability to verify reported outcomes by the responders may have influenced the conclusions drawn from the earlier studies. One prospective study using annual questionnaires, surveyed all British female medical school graduates working in hospitals during the years 1977 to 1984. Analysis showed that female anesthesiologists had no increased risk of infertility. Another study using Finnish National Health Registry data demonstrated no statistical differences between patients who had been exposed to WAG and those who had not, when medical records were examined.
However, even with the flaws in the early WAG studies, some good came out of them: waste anesthetic gas is now scavenged. A 1992 study in the New England Journal of Medicine and a later study in the American Journal of Epidemiology reported reduced fertility and increased spontaneous abortion among dental assistants employed in practices where nitrous oxide was not scavenged. And despite the fact that the modern anesthetic gases of halothane, isoflurane, enflurane, sevoflurane and desflurane are believed to be harmless in trace concentrations, their predecessors were once thought to be safe as well. Up until 1977 trichloroethylene and fluroxene were used as general anesthetics and thought to be safe. Chloroform before that. But none of the three are now in use as anesthetics because they were found to be hepatotoxic, mutagenic and carcinogenic.
The bottom line is that it’s important to scavenge waste anesthetic gases. It’s also important to stay mindful to otherwise reduce your exposure to waste gases. And finally, implement an education program for all personnel working in these areas. Studies have shown that with these procedures in place, trace anesthetic gasses can be maintained below the levels recommended by NIOSH and OSHA.
Here is a good article with more information.
In a recent clinical study, four different techniques for sealing an endotracheal tube cuff were evaluated. Eighty client-owned dogs were used in the study. Once intubated, each had its cuff inflated four times, by four different people. After each inflation, the cuff pressure was measured, the cuff deflated, and then the next technique was evaluated.
Spoiler alert: [I always jump ahead to the results.] None of the methods evaluated in this study consistently resulted in cuff pressures within a recommended range.
Each of the four anesthetists attempted one of these four techniques: (A) feeling the tension of the pilot balloon; (B) feeling the tension of the pilot balloon after a week’s practice inflating the cuff to a known pressure; (C) inflating the cuff to occlude at an airway pressure of 20 cmH2O; (D) incrementally deflating the cuff until a leak could be heard at an airway pressure of 25 cmH2O. Although the results showed none of the techniques adequate, it was encouraging to see that technique (B) approached success. It affirms the value of practice.
The article makes me wonder how effective my favorite techniques are at achieving an appropriate cuff pressure. Fortunately, it gives me the method to test them. I was also surprised at the techniques they chose to test. I was sure that by testing four different methods, I would see at least one of my favorites on the list.
The first method I learned was fast and easy. I would over-inflate the cuff, and then take my thumb off the syringe plunger and allow the pressure in the cuff to push the plunger back. When the plunger stopped moving backward, I would then add 1ml of air back into the cuff and disconnect the syringe.
In subsequent years, I learned a more precise method. It’s similar to technique (C) and (D), in that I use airway pressure to determine the cuff seal, rather than pilot balloon pressure. I seal the cuff at 15 cmH2O, but adjust so it leaks at 20 cmH2O pressure.
And now my interest in piqued. What is your favorite method for inflating an endotracheal tube cuff?The clinical study: Evaluation of the endotracheal tube cuff pressure resulting from four different methods of inflation in dogs. Vet Anaesth Analg. 2012 Sep;39(5):488-94