Fall 2016 Ten Dollars
A PUBLICATION OF SUR I N ET WOR K
Message from the President The Board of Directors of the Suri Network is pleased to bring you this 2016 issue of PurelySuri. It has been my pleasure to serve as the President of the Suri Network this year, and to work with so many outstanding and forward-looking Suri breeders. The landscape for the Suri industry– indeed, the entire alpaca industry – is constantly shifting and changing. Today, alpaca ownership is within reach of most people. New farms are getting started while some who have been in the industry for years are retiring…passing the baton to younger generations. This is the natural way of things. Soon, cria born as a result of embryo transfer will be registered in the AOA alpaca registry. While many details remain to be addressed, this is becoming a reality and we need to be involved in discussions concerning those details. A revised Suri Breed Standard – the first revision since it was developed a decade ago – has prompted excellent discussion about points that were not even at issue then, such as whether to include fiber metrics. The very fact that this discussion has occurred illuminates the progress we have made towards making a commercial Suri fiber industry a reality. Meanwhile, the Suri fiber industry moves forward. The Suri Network is undertaking a branding project this year, developing the brand for Suri fiber and products made with Suri. As your Suri breed association, the Suri Network works to provide the kind of education and services needed to preserve and promote the Suri alpaca: webinars for members, the annual Suri Symposium, PurelySuri, the Suri Herd Improvement Program, Suri rallies at alpaca shows, an online library for members, our members-only Facebook group, and active committees that address everything from fiber to the Suri Breed Standard. We believe that education empowers, and healthy discussion results in decisions that the majority supports. I am proud to belong to an organization that values all members’ ideas and opinions. I encourage all Suri owners to invest in membership in the Suri Network. This is our breed association.
Warm regards, Patty Hasselbring, President Emeritus, Suri Network
Suri Network Board of Trustees Jill McElderry-Maxwell, President - Jill began working with Suris in 2005, and purchased her first two females in 2007. She and her family moved to Maine that same year so that they could afford land for their new herd-to-be. Ten years and setting up two farms later, Bag End Suri Alpacas is now home to over fifty Suris - as well as heritage breed pigs, multiple types of poultry, Nigerian dwarf dairy goats, BLM burros, and even the occasional calf. Jill has a background in the biological sciences and paleontology, and enjoys researching and educating about alpaca health care and other topics. Jill is responsible for PurelySuri and other Network publications. Sue King, Vice President - We purchased our first Suri alpacas in 2011 after falling in love with their fine, luxurious fiber at the Oregon Flock and Fiber Festival. Our farm, Big Timber Alpacas, is located near Sherwood, Oregon and home to between 30 and 40 alpacas. We focus on breeding the highest quality Suri alpacas for seed stock. We also promote suri fiber in the commercial/cottage fiber industry, through participation in various trade shows and festivals. I have a background in corporate finance, executive leadership and general management through my career experiences with CH2M and KPMG and service on various corporate and not for profit governing boards. Deb Christner, Treasurer -My husband Doug and I raise Suri alpacas in the North Fork Valley of Western Colorado at Akuna Matada Suri Alpacas. We purchased our first alpaca in 2004 and agisted until finally, Doug and I, along with the alpacas, moved to the ranch in 2008. As chair of the Product Development Committee for three years, I was involved in creating the P2P educational DVD, the Suri Strut fashion show and educational and promotional events and publications. I have taken numerous fiber classes through the University of North Carolina, along with grading and sorting classes. I also sort and grade suri fiber for Liz Vahlkamp’s company, NASCO. Kristie Smoker, Secretary -Before starting Sweet Valley Suris, Kristie held management and executive positions in private industry for more than 20 years. She spent 18 years with the Hershey Company before co-founding Turning Point Enterprises, a human resources consultancy. Kristie currently owns and manages a farm with more than 35 Suri alpacas. She is the former President of the Mid-Atlantic Alpaca Association, a seven state Affiliate to AOA, the national alpaca association. In 2010, she coordinated the largest alpaca show in the eastern United States. Kristie received her A.S. from Pennsylvania State University and her B.S. from Albright College. Michael Alpert, Director at Large -My wife, Sherry, and I run Awesome Acres ‘Pacas & Pyrs in Oklahoma City. We have been raising and breeding Suris exclusively since 1999. I did clinical dentistry from 1975 to 1989, first in private practice and then as a Commissioned Officer in the US Public Health Service. In 1989 I was recruited by USPHS for a Systems Analyst position and served as Network Administrator and Telecommunications Manager for the Indian Health Service in Oklahoma, Kansas and Texas. I retired in 2011 and am now a full-time alpaca rancher. We also raise Great Pyrenees, and have placed dogs with alpaca ranches all around the country.
Table of Contents
Is 4-H for You?
The Suri Breed Standard
by Angie Grove
by Patty Hasselbring
Knitted Collar with Suri Locks
by Donna Rudd
Selenium Nutrition in Camelids
by Dr. Robert Van Saun, DVM
From Your Barn to Yarn: Working with Mills
by the Suri Network Product Development Committee
Don’t Pooh-pooh the Power of Poop
Mosquito-borne Diseases in Camelids
Fecal Egg Counts: Uses and Limitations
by Jill McElderry-Maxwell
by Dr. Daniela Bedenice, DVM
by Bob Storey, MS, RVT
FAMACHA© and DrenchRite©: Tools for Smart Drenching
by Bob Storey, MS, RVT
Suri Network Membership Directory
Departments 4 5 9 66
Message from the President
Board of Trustees Statement of Purpose Advertising Index
53 6 PURELYSURI
Suri Network Statement of Purpose Dedicated to the preservation of the Suri alpaca. The purpose of the Suri Network shall include, but shall not be limited to, the following: To promote, through education to the alpaca community and the general public, awareness of and interest in, Suri alpacas and their fiber, and related business interest. To promote the growth of the Suri alpaca industry. To serve as an industry and marketing group to promote and protect the collective economic and legal interests of the Network’s members. To organize and conduct, from time to time, a Suri alpaca event, which shall be open to the public and which shall further the purposes of the corporation. This event shall provide members and other participants with the opportunity to share with each other their ideas, encouragement, knowledge, and companionship.
PURELYSURI Fall 2016 • $10 PurelySuriTM magazine is a publication of the Suri Network. Statements, opinions, and points of view expressed by the writers and advertisers are their own and do not necessarily represent those of PurelySuri, members of the Suri Network, the publisher, staff, employees, or agents. Suri Network does not assume liability for products or services advertised herein. Suri Network reserves the right to accept or reject any editorial or advertising material. No part of PurelySuri may be reproduced, stored in a retrieval system, or transmitted in any form or by any means electronically, mechanically, by photocopying, recording or otherwise without the prior express written consent of the submitting author to which the article, photography, illustration, or material is copyrighted. PurelySuri assumes all work published here is original and is the work and property of the submitting author. All product and company names are trademarked or copyrighted by their respective owners. ©2016 by Suri Network, Inc. All rights reserved. Printed in the U.S.A.
Publisher: Suri Network Design & Production: Jill McElderryMaxwell
Managing Editor: Jill McElderry-Maxwell Lead Advertising Coordinator: Margit Korsak Advertising Coordinators: Jennifer Hack Contributing Authors: Angie Grove Donna Rudd Dr. Robert Van Saun The Product Development Committee Jill McElderry-Maxwell Dr. Daniela Bedenice Bob Storey Printer: Able Publishing Patient Print Guru: Steph Pride Cover Photo: Courtesy of Margit Korsak at Over Home Alpacas @2016
Photograph courtesy of Linda Kondris @2016
Suri Network, Inc. P.O. Box 1984 Estes Park, CO 80517-1984 Phone: (970) 586-5876 Fax: (970) 586-6685 firstname.lastname@example.org www.surinetwork.org
Like many parents, I was looking for an activity for my 10 year old daughter. We tried the normal gymnastic and dance classes. We even spent six weeks in a Hip-Hop class. Nothing seemed to excite her or maintain her interest. That changed in October 2010 when we visited Bent Pine Alpaca Farm’s open house. As we strolled around the farm and visited, we laughed and smiled at these new creatures. My daughter was given the opportunity to walk Sampson. The smile on her face was exciting. When we spoke to Darwin Kell and learned that he and his wife, Doris, hosted 4-H kids at their farm, my daughter was all ears. Like many, my first response was “I can’t purchase an alpaca. Where would we keep it? How would we pay for it?” Well, what is 4-H? What do you do? Do I really have time for it? Alpaca 4-H Clubs are unique, in the fact that you do not need to purchase the alpaca or live on a farm. Some farms lease you the alpaca, some assign you the alpaca for the year as long as you commit to a certain number of hours a month. 4-H alpaca clubs study everything from the anatomy, to the history, to the fiber of the alpaca. Some focus more on the fiber side and some gear more towards the livestock side. Most clubs have a monthly meeting. What 15 year old do you know that can run a business meeting and follow parliamentary procedures…well, stop by any 4-H meeting and I will introduce you. 4-H brings kids (and their families) together with others from their county and their state. While our club concentrates on alpacas, the County and State Extension offices offer camps and conferences. My daughter has made friends all across the state. What a great opportunity. In a few years when the kids are off to college, they will be making life changing decisions…where do I live, what do I want to do? How great is it to know someone in an area you may be considering to live. 4-H teaches dedication. Alpacas do not care if it is raining, snowing or 100 degrees. They need to be fed, walked and loved. Kids work with their alpaca every week. They forge a bond and trust. They learn patience, because we all know how “easy” it is to train an alpaca to walk. Have you trained yours to do the limbo? Well, if you need a hand, just call one of these kids. 4-H is a win, win for everyone. The youth learn that hard work pays off. They learn that things don’t always go as planned (ever try to walk your alpacas into the ring but they had other ideas?). Every farm owner who works with 4-H will tell you that they are proud to work with the kids, proud to teach the kids and that the kids are some of the hardest workers you will ever have. The kids can talk to the public about your alpaca, about your fiber, and about your store. In Pennsylvania, 4-H is overseen by the Penn State Extension office. Please visit their website for more information at www.extension. psu.edu. For information on the national 4-H group, visit www.4-h.org. - Angie Grove, Cumberland County 4-H Club
Is 4-H for You?
Kids and Alpacas Go Hand in Hand, and Heart to Heart
“My twins work with the alpacas in 4-H. They have gained valuable experience, knowledge and self confidence working with alpacas. 4-H fosters teamwork, friendship and citizenship in a fun learning environment. The kids love the beautiful animals and develop leadership skills that will help them grow to their full potential.” - Jacquie Gehrman, mother to Willow and Quinnie, who enjoy 4-H at Bucks County Alpacas
From www.4-H.org 4-H’ers across the nation are responding to challenges every day... 4-H is the nation’s largest youth development organization, empowering six million young people throughout the United States... Proven Results The Positive Development of Youth: Comprehensive Findings from the 4-H Study of Positive Youth Development...completed by a team of researchers at the Institute for Applied Research in Youth Development at Tufts University...shows that 4-H youth excel beyond their peers. 4-H’ers are about: • • • • •
Four times more likely to make contributions to their communities (Grades 7-12); Two times more likely to be civically active (Grades 8-12); Two times more likely to make healthier choices (Grade 7); Two times more likely to participate in Science, Engineering and Computer Technology programs during out-of-school time (Grades 10 – 12); and 4-H girls are two times more likely (Grade 10) and nearly three times more likely (Grade 12) to take part in science programs compared to girls in other out-of-school time activities.
Unparalleled Reach and Scope With 611,800 volunteers, 3,500 professionals, and more than 25 million alumni, the 4-H movement supports young people from elementary through high school with programs designed to shape future leaders and innovators. Fueled by research-driven programming, 4-H’ers engage in hands-on learning activities in the areas of science, citizenship and healthy living. Leading by Example The caring support of adult volunteers and mentors inspires young people in 4-H to work collaboratively, take the lead on their own projects and set and achieve goals with confidence. 4-H’ers chart their own course, explore important issues and define their place in the world. 4-H’ers stand up for themselves and their communities. These pivotal experiences build a foundation of leadership and skills for success in their future careers.
Suri Network Youth Program
+ Resource Book
The Suri Network has compiled a number of resources for 4-H and youth oriented activities. The PDF files can be downloaded, saved and/or printed. Members can use these documents to support local efforts with 4-H, FFA (Future Farmers of America) or other youth groups. Some alpaca farmers may also wish to lead such a group and these documents can serve as a valuable resource. Suri Network members can find the Resouce Book at http://www.surinetwork.org/Youth
Rest easy when you buy from Maine’s largest suri alpaca farm. What we can offer you... Comprehensive mentoring, before and after purchase Well trained, easy to handle animals Complete medical and fiber testing records Well conformed suris with exquisite fiber Breeding, boarding and other services available
Brion & Kristie Smoker 717-503-6168
Bag End Suri Alpacas of Maine, LLC Jill McElderry-Maxwell • (207) 660-5276 email@example.com • www.bagendsuris.com
The Suri Network Breed Standard By the members, for the members
by Patty Hasselbring, Former Suri Network President This year the Suri Network Breed Standard Committee reviewed and revised the Suri Breed Standard. It was a huge undertaking, particularly since changes in the Breed Standard must be approved by the membership. While it is impossible to achieve consensus with a broad membership such as the Suri Network’s, the Suri Network has worked hard to solicit input from both members and non-members, and to carefully consider all input in developing the current recommended draft. Certainly it would be much easier, and much more expedient, to update the breed standard with only a vote of the Board of Trustees. But, I am sure most will agree that the end product as approved by the membership is both richer and more valuable to Suri breeders. And, it is characteristic of the Suri Network to solicit input from members and to really listen to that input…it is part of who we are as an organization. We know that it is impossible to please everyone. But it is possible to hear everyone. Background The Suri Network established the first alpaca breed standard in North America ten years ago. This groundbreaking effort was tirelessly led by Suri Network member Dick Walker, with assistance and support from many other Suri Network members. A process allowing for Suri Network member input was followed by a membership vote, which approved the very first Suri Breed Standard in 2006. In early 2016 a committee was formed to review and update the Suri Breed Standard. The Board President appointed Tim Sheets and Linda Kondris as co-chairs, who then recruited members Karl Heinrich; Gail Campbell, DVM; Amanda VandenBosch; and Cheryl Gehly. Randy Coleman, Board member, served as liaison between the Board and the committee. This committee provided perspectives from alpaca judges, long-time alpaca breeders, Suri fleece experts, a fiber processor, and an alpaca owner/veterinarian. Coincidentally, the Alpaca Owners Association (AOA) identified development of a breed standard as an activity to take place in 2016. Two of the members of the Suri Network review committee are also involved in AOA’s development of the huacaya breed standard and will use the standard developed by the Suri Network for Suris as a model for the huacaya breed standard. The Suri Network News Brief in January 2016 shared the status and plans for revision, as well as the members of the committee. Process Given their mandate, the Committee went to work. They researched and reviewed the breed standards that are used by other livestock industries as well as the Australasian Alpaca Breed Standard. They identified formatting that they found beneficial. The committee used various industry resources, including The Art and Science of Alpaca Judging, published by the AOBA (now AOA); input from fleece expert Cameron Holt; and the Suri Network’s SHIP program, and of course, the original Suri Breed Standard.
Following completion of the draft revision, the committee submitted it to the Suri Network Board of Trustees. The Board of Trustees reviewed the draft and approved it for dissemination to and comment by the membership. At that time various strategies for public comment were instituted: 1.
A page was developed on the Suri Network website, with copies of the original breed standard and the proposed revisions, plus an online forum for discussion about the proposed revisions.
A letter was sent to all Suri Network members with a link to the revised Breed Standard and an invitation to review and comment.
The proposed revisions were posted on the Suri Network Membersâ€™ Facebook group.
An email was sent to all members of the Alpaca Owners Association, with a link to the proposed revisions and an invitation to comment.
The comment period was open for 30 days, allowing time for members to read and respond to the proposed revised breed standard. The Suri Network received a number of comments, both through the online forum and by email, about the proposed standards. Those comments were reviewed and discussed at length by the committee. The Result Once the comment period was completed, the committee refined the 2016 Breed Standard, addressing many of the comments that had been submitted. The final version was published for review. In addition to the final version of the Breed Standard, a side-by-side comparison between the original 2006 Suri Breed Standard and the 2016 revised Suri Breed Standard was published. Although delayed from the original timeline, vote by the membership will be held later in 2016. Ultimately, by soliciting input from members â€“ and all alpaca owners â€“ the Suri Network demonstrated once again that it is a membership organization working in the interest of all Suri owners. The proposed revised Suri Breed Standard, the side-by-side comparison between 2006 and 2016, and the original breed standard can be found at http://www.surinetwork.org/Breed-Standard. The Suri Breed Standard is developed and copyrighted by the Suri Network. The Suri Network retains its copyright on the Suri Breed Standard (both original and revised), and all rights to ownership of the document. Any use of the Suri Breed Standard must retain notice of copyright and reference to it must include credit to the Suri Network.
Photograph courtesy of Margit Korsak/Over Home Alpacas @2016
Knit Collar with Suri Locks
by Donna M. Rudd
This project is very simple and flexible. There is no rigid set of rules, rather a guide line of instructions. The lower edge is wider and longer than the top (neck edge); this is accomplished by starting at the bottom and decreasing as you knit up towards the top of the collar. The curve in the collar is made by decreasing two stitches at the middle every decrease row. No two collars are the same because the size of the yarn, needles, and locks make a difference. I prefer to use brushed Suri or Mohair yarn as it holds the slick Suri locks into the project better than regular yarn. For demonstration purposes I have used contrasting brushed yarn, you may wish to use a yarn that matches your locks. If you knit with a regular two ply yarn you should knit tighter and will likely use more locks and yarn to get the same size. I love the spontaneity of knitting to a simple pattern and adjusting things here and there. First you will need a supply of washed locks. I wash locks by first separating them and placing them side by side in wash bags where they are washed and rinsed before they are laid out to dry. Before knitting any excess debris or stray fibers are removed from the locks as I like the locks to look crisp and tidy all the time. This is one reason I knit from the bottom of the project to the top and add the locks to the outside (facing away from me) as I am knitting the rows. I keep a slick smooth scarf on my lap when knitting so that the locks do not become roughed up while knitting. Spritzing the project with water occasionally will settle static and help the locks keep their shape. I used # 4 mm/ 6 American double pointed 24 inches long needles, but you can use whatever you like to get a gauge to roughly four stitches per inch (this will vary somewhat depending on the size of your locks and yarn). Donna Rudd is a Master Spinner and long time Suri fiber enthusiast living in Canada.
1. Cast on 150 stitches to obtain a length of approximately 30 inches (this is the bottom row of the collar and the longest portion) 2. Row 1-3; Knit 3. Row 4; * k3; next 3 stitches ( #1, insert lock with yarn (tip of lock facing right and cut end knit into stitch with yarn); #2, knit stitch and include last of cut end of lock if it is protruding; #3 stitch k1;) k3* repeat from * to * to end of row …… end with k 3. See following illustrations.
4. Row 5; Knit 5. Row 6; Knit 2, K2 together, knit to middle of row, knit 2 together, continue knitting to last 4 stitches, knit 2 together, knit 2 6. Row 7; Knit 7. Repeat rows 4-7 until project is size you wish then bind off after row #7 (this is the top of the collar, the narrowest part) 8. Cast Off; * K1, K2 together*, slide first stitch over second stitch to drop one, repeat until all stitches are cast off. You may wish to add a yarn loop at the end of the bind off row and a button on the opposite side. If you prefer a ‘fuller’ collar with more locks, then on the row where you add the locks add them every second stitch instead of every third stitch.
Long and short edges
Row 4 (adding locks)
Selenium Nutrition in Camelids Dr. Robert Van Saun, DVM, MS, PhD
Selenium (Se) is one of a number of essential microminerals required by llamas and alpacas as well as all other animals and people to help maintain normal body functions. Selenium is termed a micromineral, in contrast to a macromineral like calcium, because it is required in very small quantities; milligrams (mg) per day. However, Se has a notorious history that places it in a unique situation relative to feeding recommendations and regulations. The objective of this article is to provide an overview of Se nutrition relative to biologic functions and associated disease conditions, as well as to address appropriate supplementation practices, evaluating mineral products, and monitoring Se status in an effort to keep llamas and alpacas healthy.
Selenium was not determined to be an essential mineral until the late 1950â€™s. Immediately following this discovery, Se supplementation was observed to prevent a disease process termed â€œstiff lamb diseaseâ€? or what has become known as white muscle disease or nutritional myodegeneration. It was not until 1973 when Se was determined to be a functional component in the cellular antioxidant, glutathione peroxidase (GSH-Px). A number of essential and nonessential nutrients have biologic roles as antioxidants that collectively protect cell function and structure. There is tremendous interest in dietary antioxidants in human and animal nutrition as they have been linked to preventing or protecting against heart disease, aging, cancer, and many other diseases. All cells undergoing normal metabolism generate potentially toxic end products termed reactive oxygen species (peroxide radicals) that have strong oxidizing capabilities. Oxidizing metabolic byproducts, when left unchecked, can result in damage to cellular components including chromosomes, proteins, and cell membranes ultimately destroying the cell. Selenium is only one of a number of biologic antioxidant agents the body has at its disposal to inhibit damage from internal or external (e.g., pollution, UV radiation, smoke) oxidizing agents. Another well known biologic antioxidant is vitamin E, which works in concert with Se to collectively protect cell membranes (vitamin E) and cell contents (Se) from oxidative damage. The interrelated antioxidant function of vitamin E and Se accounts for why they are often found together in nutritional supplements. Additional dietary vitamin E or Se can replace the other in situations where one is marginally deficient. Other biologic functions of Se are not fully understood, though a limited number of selenoproteins have been identified. More recently it has been discovered that one of these selenoproteins plays an important role in thyroid function. Secretions from the thyroid gland are important regulators of cellular activity and overall body metabolic rate. This deiodinase selenoprotein converts thyroxine (T4) to the biologically active form triiodothyronine (T3), which mediates rate of cellular metabolism. In the presence of Se deficiency, T4 can accumulate while T3 levels will diminish, thus inducing a state of hypothyroidism.
A wide spectrum of disease conditions have been attributed to Se deficiency, though not all have been well documented to be true deficiency conditions. White muscle disease (nutritional myodegeneration) is the best recognized clinical Se deficiency disease. Any age animal can be affected, though younger animals most commonly experience clinical disease. As the disease name implies, severe Se deficiency results in pathologic degeneration of skeletal muscle fibers with secondary fibrosis (Figure 1).
Figure 1. Microscopic photograph of skeletal muscle fibers undergoing degenerative changes associated with clinical selenium deficiency. Most fibers are affected and showing signs of swelling and loss of striations typical of normal fibers (long arrow). Some muscle fibers are being replaced with fibrous tissue (short arrow).
Figure 2. Heart from a calf with white muscle disease. Notice the pale areas of the heart wall muscle (Photograph courtesy of Dr. John King, Cornell University)
These lesions change the physical appearance of muscle tissue from its normal red to a pale white color. Affected clinical animals will show signs reflective of specific muscles affected and severity of degenerative changes to muscle fibers. Typically both hind legs are symmetrically affected; however, tongue and heart muscles are commonly involved in newborn or young growing animals (Figure 2). With skeletal muscle damage, affected young or older animals will show various degrees of lameness, weakness, or difficulty moving. Acute death can occur in those younger animals where the heart muscle is damaged. Newborn animals with tongue lesions will have difficulty nursing and may be diagnosed as a “dummy” animal. Severe Se deficiency has been attributed to causing abortion and stillbirth. All of these problems have been documented in most domesticated species and believed to occur equally in llamas and alpacas. Selenium also influences immune cell function and marginal deficiencies will result in an increased susceptibility to disease. Most studies have shown a critical role for Se in the nonspecific immune response. Cells that engulf bacteria (phagocytes) are unable to kill the ingested bacteria when the animal is Se deficient. Other studies have suggested that Se also influences the body’s ability to mount an appropriate antibody response to an infectious agent. Subclinical Se deficiency in growing animals, through its affect on immune response, may predispose them to diarrhea and pneumonia conditions. Young animals infected with coccidia may not be able to mount a sufficient immune response to help them recover from the disease. This will result in a prolonged disease condition and a perception of disease treatment failure. Adult females with marginal Se deficiency may be more susceptible to uterine infections (metritis) around the time of breeding. Premature, weak, or poor doing babies have been attributed to Se deficiency. Although there are many potential causes of “ill-thrift” babies, selenium’s role in thyroid function might explain a possible link to this disease syndrome. Deficiency problems primarily gain our attention when discussing Se, but we should not forget that Se is more notorious for its toxicity concerns. Consumption of specific Se-accumulator plants can result in an acute Se toxicity that occurs over a period of hours to days. More common is a chronic Se toxicity syndrome termed alkali disease. The disease is associated with prolonged consumption of seleniferous plants. These plants, and the high Se soils on which they grow, are scattered throughout the northern Great Plains of North America. This disease was first recognized in the Dakota’s and Nebraska during the 1860’s and has even been suggested to have contributed to the defeat of General George Custer at the Battle of Little Bighorn. Alkali disease is characterized by cracks and lesions of the hoof wall, abnormal hoof wall growth, brittle hair, and hair loss. Affected animals are often in poor body condition (emaciated) and show various degrees of lameness. Of greater concern to llamas and alpacas is acute Se toxicity. With the greater propensity for Se deficiency in many regions where llamas and alpacas are raised, more owners are concerned with supplementing Se. One mode of Se supplementation is to inject a commercial Se product. One must be careful in using injectable Se products as their concentration varies, thus the dosing amount will vary (Table 1). Suggested dosage will range from 1 ml per 40 lbs (Bo-Se®) to 1 ml per 200 lbs (Mu-Se®) of body weight. With injectable Se one wants to be very careful in the amount given as over dosage can result in acute toxicity. Injectable Se has high biologic availability and is readily absorbed. There is no antidote for a situation of acute Se toxicity. In such cases the animal will show signs of distressed breathing, salivation, and cardiovascular collapse. This all may occur within minutes to an hour following an injection of an excessive amount of Se. Toxic dosages have not been well defined for all species, but more than 0.5 mg/kg body weight is considered toxic for sodium selenite injections. This is about 20 times the suggested dosage for these products (Table 1), but easily achieved by using an inappropriate product. Acute Se toxicity needs to be differentiated from an anaphylactic reaction, which can occur with Se injections. Anaphylactic reactions can be successfully treated with an appropriate dosage of epinephrine.
Table 1. Comparison of selenium content and suggested dosage for injectable selenium (sodium selenite) products. Selenium Product1
E-Se® Mu-Se®, Velenium®
2.5 mg/ml 5 mg/ml
Recommended Selenium Dosage 25 µg/lb body weight 25 µg/lb body weight 25 µg/lb body weight 25 µg/lb body weight
Dosage Amount 1 ml per 10 lbs body weight 1 ml per 40 lbs body weight 1 ml per 100 lbs body weight 1 ml per 200 lbs body weight
L-, Bo-, E- and Mu-Se products from Shering-Plough, Velenium product from Fort Dodge. All are prescription products only available from veterinarians. Not currently being manufactured.
A number of veterinary diagnostic laboratories are finding very high concentrations of Se in llama and alpaca liver samples, suggesting an excessive level of supplementation. Some laboratories have identified Se toxicosis as a potential contributor to the death of the animal in a number of these cases. Whether the high liver Se concentration is due to injection or oral Se supplementation has not been clearly defined. Our understanding of Se metabolism in llamas and alpacas is very limited, especially related to injectable Se distribution, and requires further research. In some of these “toxicity” cases, no injectable Se supplementation was documented, suggesting excessive oral supplementation.
Due to toxicity concerns as previously discussed, addition of Se to animal diets is regulated as a food additive by the Food and Drug Administration. It was not until 1978, some 20 years after the discovery of Se essentiality that animal feed manufacturers were permitted to include supplemental Se in animal feed products. Being classified as a food additive, this means Se cannot be used as a medicine and veterinarians are not able to write prescriptions for higher level of use. Addition of Se was initially allowed for poultry and pigs at an incorporation rate of 0.1 parts per million (ppm) of the total diet. Over the next couple of years, this allowance for Se inclusion was expanded to include cattle, sheep, and goats. Field experience with dietary Se supplementation was suggesting that the allowable level of 0.1 ppm was not sufficient to prevent deficiency disease problems. After considering the available scientific information, in 1987 the FDA increased the allowable supplemental dietary Se incorporation rate to 0.3 ppm of the total diet. Use of organic Se (selenomethionine) from Se-fortified yeast cultures was approved in 2000 for chickens and then other species over the next couple of years. Llamas and alpacas are not mentioned in any of these FDA regulations. By the letter of the law, this would mean supplemental Se cannot be legally added to their diets. However, this is not a situation the FDA would pursue as long as the current regulations for similar species are being followed. Current FDA regulations allow supplemental Se to be added to the total diet at a level of 0.3 ppm. However, Se cannot be easily added to forage, thus making it difficult to supplement Se for animals consuming primarily forage-based diets. To address this issue, the FDA permits Se to be incorporated into mineral mixes for animals. Again, the Se concentration of a mineral mix is regulated to conform to expected total dietary intake of Se from the product. Mineral products formulated for sheep and goats are allowed to have a maximum of 90 ppm Se, while products for cattle can contain a maximum of 120 ppm Se. These Se concentrations are based on estimated mineral intake and the total amount of Se to be consumed to meet the 0.3 ppm of supplemental Se in the total diet.
Selenium in its crystalline form from the Anna Mine, Alsdorf, Aachen, North Rhine-Westphalia, Germany http://www.mindat.org/min-3611.html (Photograph courtesy of Frank de Wit, www.mindat.org)
For example, beef cattle are estimated to consume on average about 10 kg (22 lbs) of a forage diet per day. If one is to supplement Se at the legal level (0.3 ppm), then a beef cow could consume 3 mg Se per day (0.3 mg/kg x 10 kg/day = 3 mg/day). If this 3 mg is to be packaged into a mineral product, the Se concentration of the mineral will need to be adjusted to expected mineral intake. For beef cows this is estimated between 0.75 and 1 oz per day. In completing the calculations to package 3 mg Se into 0.75 oz, this works out to 120 ppm Se in the salt (refer to Table 1A). Similar calculations were undertaken for sheep and goats on the basis of meeting the defined supplemental Se amount of 0.7 mg/day. Based on a lower expected mineral intake for sheep and goats, allowed maximal Se content of free choice mineral is 90 ppm Se. In allowing the use of Se-fortified mineral supplements, the expectation is the mineral will be the only source of supplemental Se in the animal’s diet. To make this issue even more confusing, the allowable Se content for mineral mixes is a maximal value. Feed manufacturers can add any level of Se to their mineral products up to the maximal value for the given species of animal. Therefore, you will need to assess the feed tag information for your mineral product to determine Se content and expected intake rate. Compare expected intake to your animal’s mineral intake to determine if the Se content is appropriate to meet daily need (use Table 2A for comparisons). Table 2. Calculated amounts of either selenium intake from free-choice mineral supplements (A) or selenium-fortified pellet supplement (B) needed to achieve specified levels of selenium intake. A. Free Choice Mineral Supplements Daily Salt Intake (oz) Selenium Mineral Content 0.25 0.33 0.5 (ppm) Amount of Selenium (mg/day) consumed 30 0.21 0.28 0.4 50 0.35 0.47 0.7 70 0.50 0.66 1.0 90 0.64 0.84 1.3 120 0.85 1.12 1.7 B. Selenium Containing Pellet or Grain Supplements Supplement Selenium Concentration (ppm) Desired Selenium Intake 0.3 1.0 2.0 mg/day lbs supplement needed to be consumed per day 0.5 3.7 1.1 0.6 0.75 5.5 1.7 0.8 1.0 7.3 2.2 1.1 1.5 11.0 3.3 1.7 2.0 14.7 4.4 2.2 2.5 18.4 5.5 2.8
0.85 1.4 2.0 2.6 3.4
1.1 1.8 2.5 3.2 4.25
0.3 0.4 0.55 0.85 1.1 1.4
0.15 0.20 0.27 0.43 0..55 0.7
Reports from the National Research Council (NRC), a scientific body that reviews available research to determine nutrient requirements for animals, consider the dietary requirement for Se to be between 0.1 and 0.3 mg/kg of dietary dry matter (DM). This requirement range was defined for all production animal species from pigs and horses to sheep and cattle. A new NRC report on small ruminant nutrient requirements (http://books.nap.edu/catalog.php?record_id=11654), which includes llamas and alpacas, recommends 0.74 mg Se/ day, or 0.2 mg/kg of diet DM for llamas and alpacas. Unfortunately there are no feeding trial reports to document this requirement. This NRC recommendation is a minimal value and based on extrapolations from other species and information from a single published survey of Se supplementation in llamas (Herdt, 1995). In this survey, supplementation around 1 mg Se/day to adult llamas was associated with adequate blood Se concentrations and ability to maintain normal Se concentrations in crias born to these females. Another fundamental challenge in defining Se requirement for camelids is the documented difference in dry matter intake compared to other species. Llamas and alpacas consume less food per unit of body weight compared to sheep, goats, and cattle. This results in a disconnection between dietary requirements and conforming to current FDA regulations. In using the current daily (0.74 mg/day) and dietary content (0.2 mg/ kg) recommendations, predicted total intake would be 3.7 kg (0.74 mg/day ÷ 0.2 mg/kg = 3.7 kg/day). This intake greatly exceeds any estimated intake for llamas in the NRC report. These differences underscore the need for further research on mineral requirements for llamas and alpacas. Dietary Se content issues must be resolved; however, the recommended daily amounts of Se to supplement, between 0.74 and 1.0 mg/day, are reasonable guidelines to incorporate into one’s feeding program.
To this point I have discussed supplemental Se in the diet. Inherently there is some Se content to feed ingredients. With llamas and alpacas, the primary feed ingredient is forage with supplemental concentrate (pellets, mineral, or both). Forage Se content is extremely variable across all of North America and dependent upon soil conditions. Selenium content of the soil is variable (ranging from < 0.1 to > 80 ppm) and soil acidity, rainfall amount, and other factors can greatly influence its availability to plant tissues. Acid soil conditions, heavy rainfall, and presence of inhibiting substances (iron and aluminum) will result in very low plant Se content. Essentially the eastern coast, north to south, over to the Great Lakes region and the entire western coast areas are low (< 0.1 ppm) in forage Se content. Most all Canadian provinces are low in Se. Only the central plains states up into Manitoba and Saskatchewan have moderate to high soil Se and variable to high plant Se content. Although difficult, forage Se content can be determined at some forage testing laboratories. Unless you have forages from these Se-adequate areas, you should ignore the Se contribution from forage and add the maximal amount of supplemental Se via mineral or pellet products. As previously described, free choice mineral products can range widely in Se content; from minimal (10 ppm) up to the legal maximum. What you are interested in is the total amount consumed. To assess mineral adequacy relative to Se, one needs to determine the Se content of the mineral and daily animal intake. Mineral Se content can be determined from the feed tag with the product. Selenium content may be expressed as ppm (mg/kg) or as a percent (%). To convert percent to ppm, move the decimal point to the right 4 places. For example, 0.005% Se is the same as 50 ppm. The bigger challenge is determining average daily mineral intake. Mineral intake is controlled by salt content of the mineral product. Most products will have some intake guidelines on their feed tag. However, expected intake is often over estimated. Mineral intake will be variable over time, but typically llamas and alpacas can be expected to consume between 0.25 and 0.33 ounces per day. Again, it is best to determine this for your animals. Also, do not have both white salt and a trace mineral salt available for the animals to choose. They only seek out a salt source. Armed with intake and Se content information, you can then use Table 1A to assess Se adequacy. In reviewing this table, the highlighted cells show the combination of mineral Se content and intake that achieves at least 1 mg Se intake per day. From these data it can be seen that only mineral that has at least 90 ppm Se will achieve near 1.0 mg Se intake with a daily mineral intake less than 0.5 oz per day. Many of the commercial mineral products contain less than 90 ppm Se.
Alpacas grazing at La Raya Research Station at 14,000â€™ elevation. Soils in the altiplano are naturally much higher in selenium content than those in North America (photograph courtesy of Dr. D. Andrew Merriwether)
Another method of supplementing Se is through the pellet or grain supplement. Using the data shown in Table 2B, one can determine a reasonable Se concentration for their pellet or grain product. If we set 1.0 mg Se/day as the goal (highlighted row in Table 2B), read across the row to see how many pounds of concentrate would need to be fed to achieve this Se intake amount. The variation is due to the different concentration of Se in the pellet or concentrate product. In this table the Se concentration of the pellet or concentrate is varied from 0.3 to 8 ppm. It is only when you have a Se concentration of 2.0 ppm or greater where you would be feeding 1 lb or less of the pellet or concentrate product to achieve the desired 1.0 mg Se/day delivery rate. These two examples demonstrated how one could provide the entire supplemental Se allotment from either mineral or pellet sources. Be careful not to provide both sources and potentially double the amount of supplemental Se. We have highlighted potential problems with Se toxicity, is there a concern with oral supplementation? Nonruminant animals such as pigs and horses absorb Se from the diet very efficiently and hence are susceptible to toxicity problems. The maximal tolerable level for Se in the total diet of nonruminants is 2 ppm or about 10x the requirement. Remember, this is the total diet consumed and not a single ingredient. In contrast, ruminant animals, including llamas and alpacas, are less efficient at Se absorption due to rumen alteration of the Se molecule. Though not determined directly for llamas and alpacas, maximum tolerable level for Se in ruminant diets is considered 5 ppm or even higher. This means that ruminant animals, including llamas and alpacas, are less susceptible to Se toxicity, but with excessive supplementation it can happen. There are still many questions to be answered relative to Se supplementation in llamas and alpacas.
Monitoring Se Status
One can do their best to provide a balanced diet that contains sufficient Se to meet current dietary recommendations. However, we cannot ensure that the animals consume the diet to be specifications we desire. To this end we should incorporate some method of evaluating nutritional status monitoring process to ensure neither deficiency or toxicity potential exists. Selenium content of the diet is difficult to determine and an expensive procedure. One is best served by monitoring Se content of supplemental feeds (mineral or pellets) and determine response of the animals. Fortunately, Se is one of the nutrients that can be adequately assessed through the use of blood concentrations, though there is some debate on the preferred method. Selenium concentration can be determined in serum (without cells), whole blood (with cells), or in tissue (primarily liver) as a method of Se status assessment. Selenium status can also be assessed by determining glutathione peroxidase (GSH-Px) activity in blood. Laboratories that offer GSH-Px analysis are limited and it is a more expensive and technically difficult procedure. Liver Se content is a good determinate of Se status, but one must obtain a liver tissue sample either by biopsy on a live animal or from a dead animal. It is well worth the cost to have liver mineral analyses completed on any animal that dies (young or old) as a routine monitor of nutritional status. Most laboratories will perform either serum or whole blood Se concentration analyses. Serum Se concentration reflects more acute or recent changes in Se nutrition, whereas whole blood Se reflects more chronic or historical Se status. This is a result of GSH-Px residing primarily in the red blood cell and each red blood cell lives for 105 or more days. However, llamas and alpacas are slightly different from other species. They have more GSH-Px outside the red blood cell and thus have generally higher serum Se concentrations compared to other species (Table 2). Table 2. Diagnostic criteria for evaluating serum selenium concentrations (ng/ml) in sheep, goats, and camelids (data based on Michigan State Nutrition Laboratory values). Age Category
Neonate (1-9 days)
45 - 80
75 - 130
Suckling (10-29 days)
50 – 90
75 - 130
Weanling (30-150 days)
60 – 90
75 – 150
Yearling (151-500 days)
70 – 110
130 – 230
100 - 145
130 - 230
Many laboratories do not have extensive databases to determine appropriate reference values for llamas and alpacas. Therefore, interpretation from a given laboratory might vary given the reference values being used. Selenium concentrations in serum and whole blood will also vary by age of the animal. These are important considerations if one is to properly interpret laboratory results. These observations may partially explain some of the current issues of interpreting high (toxic) Se concentrations in llamas and alpacas that have not been known to be exposed to toxic intake or injections. References Herdt, T.H. Blood serum concentrations of selenium in female llamas (Lama glama) in relationship to feeding practices, region of the United States, reproductive stage, and health of the offspring. J. Anim. Sci. 1995;73:337. National Research Council. Nutrient Requirements of Small Ruminants, National Academy Press: Washington, DC, 2007. Van Saun, R.J. Nutrient requirements of South American camelids: A factorial approach. Small Ruminant Research 2006;61(2/3):165. Van Saun, R.J. Feeding the Alpaca (Vicugna pacos), In: Hoffman, E., ed., The Complete Alpaca Book, 2nd ed., Bonny Doon Press: Santa Cruz, CA, 2006.
Dr. Robert Van Saun is a distinguished Professor of Veterinary Science at Pennsylvania State University. He specializes in nutrition, metabolic disease, and nutrition/reproduction interactions in large and small ruminants, particularly South American camelids. PhD, Ruminant Nutrition, Department of Animal Science, Cornell University, 1993 MS, Large Animal Clinical Sciences, College of Veterinary Medicine, Michigan State University, 1988 Residency, Theriogenology, Field Service Section, College of Veterinary Medicine, Michigan State University, 1987 DVM, Veterinary Medicine, College of Veterinary Medicine, Michigan State University, 1982 BS, Zoology, Michigan State University, 1978
From Your Barn to Yarn: Working with Mills Selected Excerpts from the Resource Directory-All things Suri, Prepared by the Product Development Committee. http://surinetwork.org/resources/Documents/Fiber%20Resource%20Guide%202016.pdf Educational DVD, P2P-Disc 3, Chapter 2 There are two main systems of preparing fiber for yarn: the worsted system and the woolen system. The worsted system is defined by the removal of short fibers by combing and the preparation of top by gilling. In the woolen system, short fibers are retained, and the process may or may not involve combing. The diagram below shows the steps involved in both systems: the blue lines and boxes show woolen processing, and the green lines and boxes show the steps in worsted processing. Final products produced before the spinning stage are shown in the yellow boxes to the left. The chart on the facing page shows what products are produced at each step in the processing chain, with final products shown by the machine that produces them.
GRADED FLEECE WOOLEN PROCESSING
FIBER SEPARATOR (DEHAIRER) PRODUCTS ROVING, BUMPS BATTS, BATTING CARDER
DRAFTING-DRAW OR PIN
SPINNING & PLYING
Raw Fiber Fiber ready for shipment to mill
“The Fiber Washer”
Cloud of prime & waste fibers
Needle or Wet Felting
“Core Spun Attachment”
“Comb Top Machine”
“Pin Drafter or Draw Frame”
Single ply yarn
“The Draw Frame”
“The Plyer” “The Steamer”
2,3,4 or novelty yarn product
“The Cone Winder”
Cone of yarn
“The Skein Winder”
Skein of yarn
Ball of yarn
TUMBLING Before tumbling any fiber, always check with your processor, some do not want you to tumble your fleeces. The tumbler is a mechanical skirting aide. It tumbles the fiber, allowing dirt, vegetation, short fibers, and guard hair to be released from the rest of the fleece and fall to the floor. The tumbler can be used either before or after washing. It can also be a big asset to fiber producers, as tumbling your fleeces on your farm can save you money in shipping and add quality to the end product.
SCOURING-WASHING The fiber wash system will wash a large amount of fiber in a single operation. Up to three lots of fiber can be cleaned at the same time with variable wash action and centrifuge. The integrated recycling system allows reuse of approximately 40% of the water and 30% of the heat.
PICKER The picker opens and blends fibers. Picking improves homogeneity and is the best stage to blend colors or fiber types. Opening reduces fiber entanglement. Entanglement in natural fiber occurs on the animal from exposure to weather or rubbing. It can also be caused by washing. At the picker, fibers are fed at low speed onto a rotating drum. The fibers contact pins on the drum, and are rapidly teased out. Given the drum speed, the fibers are thrown from the rear of the machine into a “collecting room”. Most of the opening is accomplished by air movement caused by the speed of rotation and the vanes on the drum. Mixing fibers is called blending. There are noticeable differences in the fiber within a fleece and between fleeces from animals of the same breed. These differences can be evened out so that yarn is identical in characteristics from the start to the end of the batch.
FIBER SEPARATOR (DEHAIRER) Also known as a “dehairer”, the fiber separator is deigned to accept clean fiber and selectively separate finer fiber from coarser fibers and vegetable matter. Unwanted materials are collected in a chamber beneath the machine, and finer fibers exit the machine at the end in the form of a web.
CARDER The Carder is the heart of the fiber processing mill. All fiber must be carded successfully. We may define the carding process as the conversion of a random mass of fiber into a continuous web, either to form a “batt” for felt making, or drawn together to form a “sliver”. However, this simple definition does not convey the complexity of the working with the fibers, which is of fundamental importance. Individual fibers must be separated from each other, aligned, and delivered from the carder in a consistently even web.
CORE SPUN ATTACHMENT With this attachment we can create a “core yarn” at the carding stage. A core yarn is simply a core material wrapped tightly with fiber. The core ensures durability and longevity and gives mats and rugs the required weight to lie on the floor. This machine is capable of making quality rugcraft yarn from lower grades of fiber. Fiber artists are knitting, crocheting, braiding, and weaving with these yarns. It is also capable of creating coreless lopi yarns.
COMBING Combing is a method for preparing carded fiber for spinning. Only a few combing machines exist in the United States. Combing is divided into linear and circular combing. The Noble comb is an example of circular combing. The French comb is an example of linear combing. The process of combing is accompanied by gilling, a process of evening out carded or combed top making it suitable for spinning. Combing separates out short fibers by means of a rotating ring or rectilinear row of steel pins. The fibers in the “top” it produces, have been straightened and lie parallel to each other. Pictured is a combined gilling box and combing machine.
DRAW FRAME Drawing or drafting is the reduction in weight relative to length or linear density of the fiber stream or sliver. It is achieved by passing the sliver through the nip points of two sets of rolls: a lower steel roll and an upper rubber coated roll or “cot”. The front set of rolls has a higher surface speed than the rear. The sliver is elongated (or drawn) by a factor equivalent to the ratio of the speeds of the two sets of rolls. To manage fibers of varying length we use a porcupine roller that also improves the parallelization of the fibers. The operation of drawing sliver is quite straightforward and simple in comparison to the complexity of carding, but it is important as a means of establishing the linear density of the sliver to a level suitable for spinning.
PIN DRAFTER The pin drafter is used to further align the fibers in the sliver after the card. This has several effects on the sliver. First, it aligns the fibers to a more parallel state and second, it takes several slivers at once and blends them together. This produces one sliver out that is more even than the slivers produced by the card. Fibers may be pin drafted to align them further into a parallel sliver. This allows an easier draft and produces a “semi-worsted” type yarn. Fibers must be pindrafted several times to prepare them for machine spinning.
SPINNER Spinning is the process by which twist is inserted into a sliver. There are two parts to this spinning system, the draft and the twist. The ring spinning frame has a wide range of possible rates of draft and twist insertion. Depending on fiber fineness and fiber length, a very wide range of yarn types may be produced. A “master dial” adjusts the speed of the front and back rollers and the spindle speed. By correct adjustment of the control knobs, a yarn of the desired count (thickness) and twist level may be produced. The spindle direction may be reversed to give “S” or “Z” twist direction and to allow twist reversal, for the purpose of plying the single yarns. The required amount of twist inserted into a yarn depends upon the count and the type of fiber.
PLYER Plying is performed on a four spindle spinner by bypassing the draft zone and reversing the direction of twist. There are many aesthetic changes created by plying single yarns. The most obvious is the development of loft and softness. Plying single yarns of different count, twist level, and color may produce fascinating results. In single yarns, twist gives the fibers cohesion and strength. However, a single yarn can be hard and unsuitable where softness and warmth are important. To achieve loft and softness the yarn must be twisted in the opposite direction to that in which it was spun, with two, three, four or five single yarn strands. A great deal of variation in plied yarn properties may be achieved by varying the amount of twist inserted into the plied structure.
STEAMER This piece of equipment is one way to “set” the yarn. Yarn is passed through a chamber where it is exposed to live steam under slight tension. It is then allowed to cool as it passes through a stainless steel tube.
CONE WINDER Yarn is removed from the bobbin and wound around a cardboard cone. This allows for larger amounts of continuous yarn to be removed from the bobbin. Many times weavers prefer cones to skeins.
SKEIN WINDER A skein is a measured length of yarn which has been wound into a large circle and then is usually twisted into a convenient shape for sale, storage or further processing such as dyeing. The winding operation is essentially the removal of single or plied yarn from the bobbin into a reel. This time consuming manual task is accomplished at high speed producing several skeins simultaneously.
What you should know about your mill and some questions to ask... • Who are they? Are they a large or small organization? The size of a mill may affect turn around time, communication and many other facets of processing. • Where are they? Will you need to pay for shipping your fiber? • What are their hours? Can you call at all times? • What is their pricing structure? Some mills charge by incoming weight, others by outgoing. The difference in price may be significant. • What equipment do they have? Can they make the kind of products you’re interested in? Not all mills can make felt or corespun yarn, for example. • What is included in each pricing group? For example, when asking about yarn, what steps are included? Washing, picking, carding, spinning? • How do they feel about processing Suri? Not all mills are created equal. Ask other Suri owners for their recommendations. • Will they make it “to order”? • Do you have a choice of skeins or cones? Who are your customers, and which will they want? • Do they make recommendations for your fleece? This may be especially helpful for those new to processing fleece. • Can they supply blending fibers should you desire? • What is expected delivery time? Turn around time varies by mill and often by time of year. • Can your order be expedited? • What additional services do they provide? • How do they want you to pay? Is a deposit required? • Are they open to tours andor visits? • Do they attend events where you could drop off your fleece?
2016 Fiber Mills that Process Suri Fiber This is a listing of fiber mills that have indicated to the Suri Network that they are interested in processing Suri fiber. No recommendation of any individual mill is expressed or implied. A&B Fiberworks Arizona Fiber Mill Anne Goodwin Rita Beeson PO Box 678 7485 E 1st St Ste A Unden Canada TOM1JO Prescott Valley, AZ 86314 403-443-5907 928-713-3244 firstname.lastname@example.org Rita@arizonafibermill.com
Blue Hills Fiber Mill Ridgeley Reichert W12855 Christianson Rd Bruce, WI 54819 715-868-9045 email@example.com
C & M Acres Fiber Mill Christian Davies 33707 663rd Ave Maxwell, IA 50161 515-387-8607 firstname.lastname@example.org
Central Virginia Fiber Mill Mary Kearney 1710 Welsh Run Rd Ruchersville, VA 22968 434-985-3669 CentralVAFiberMill@gmail.com
Crooked Fence Alpacas & Mill Linda Kenison HC 60 Box 85 Mona, UT 94645 801-367-1629 email@example.com
Dakota Fiber Mill Chris Armbrust 17061 54th St, SE Kindred, ND 58051 701-238-4002 DakotaFiberMill@gmail.com
Fitch Fibers Linda Adelman 25 Stockhouse Rd. Bozrah, CT 06334 860-222-3119 FitchFiber@gmail.com
Going to the Sun Fiber Mill Diana Blair 805 Kienas Rd Kalispell, MT 59901 406-756-6772 MontanaMill@gmail.com
Luxe Fiber Designs Linsey & Matt Carey 11282W Murphy Blvd Hayward, WI 54843 715-214-5963 Alpacaroyale@hotmail.com
Morro Fleece Works Sheri McKelvy 1920 Main St Morro Bay, CA 93442 805-772-9665 firstname.lastname@example.org
Mystic Pines Fiber Processing Rob Jorissen 7892 N Trails End Dr Williams, AZ 86046 480-326-7279 Info@mysticpinesfiber.com
New Aim Fiber Mill Nancy Williams 13 Robinson Rd Waldoboro, ME 04572 207-832-5162 email@example.com
New Era Fiber Jan Heinrich 698 Wallace Rd Gallatin, TN 37066 615-452-7852 firstname.lastname@example.org
Rach-Al-Paca Fiber Processing Rachel Boucher 839 Third St Hastings, MN 55033 651-485-7916 email@example.com
Ranch of the Oaks Tom Goehring 18495 Goodwin Ave Lampac, CA 93436 805-714-2068 Ranch@ranchoftheoaks.com
The Shepherdâ€™s Mill, Inc. Jay & Sally 3269 Crucero Rd Phillipsburg, KS 67661 785-543-3128 firstname.lastname@example.org
Spring Harvest Fiber Mill Denise & Bob Cathel 2361 Scoon Rd Sunnyside, WA 98944 509-837-8012 email@example.com
Sterling WoolNook & Mill Deidre Bushnell 1327 Byerland Church Rd Willow St., PA 17584 717-371-4195 firstname.lastname@example.org
Still River Mill, LLC
Star Castle Fiber Mill & Farm, LLC Barbara Simpson 56778 Fat Elk Rd Coquille, OR 97423 541-400-0081 Barbara@starcastlefarm.com
Sugarloaf Alpaca Company Nancy Brandt 1347 Buckystown Pike Adamtown, MD 21710 301-606-2133 Nancy@sugarloafalpacas.com
Vermont Fiber Mill & Studio Ed & Debbie Bratton 185 Adams Rd Brandon, VT 05733 802-236-9158 email@example.com
210 Eastford Rd Eastford, CT 06242 860-9974-9918 firstname.lastname@example.org
Don’t Pooh-pooh the Power of
Poop... Make Manure an Alternate Income Stream for Your Farm by Jill McElderry-Maxwell
Let’s face it - beans don’t get much respect. We spend so much time sweeping them, raking them and putting them in piles, but how many of us actually use them? Hands up if you put your composted manure back on your fields, or use it in your own gardens. That’s great! You know just how much plants love paca poop - they grow taller, thicker, and more vibrantly, producing more flowers, fruits and vegetables than plants grown in plain soil. You probably have the best tomatoes on the block. Alpaca manure doesn’t burn plants even when fresh, and when aged, it’s a good source of organic nitrogen, trace minerals, and carbon. It adds tilth to your soil, benefitting both the macro and microbiota, from red wigglers to beneficial nematodes. Now, put your hands up if you make money from your manure. Every farm needs any income stream it can find, and for some of us, that includes cashing in on an abundance of beans. Several national publications published articles on the value of alpaca manure this spring, and at Bag End at least, the phone has been ringing off the hook with people looking for poop. I had a couple drive two hours each way to my farm for a trailer full of manure! I’m also going to be enjoying vegetables from a startup CSA that hauled away a whole heap of my manure pile. While I will happily load manure with my tractor, other farms offer the option to selfscoop, or sell prefilled grain sacks or buckets of manure. A simple Craigslist ad can guarantee that your bean pile will vanish, leaving behind a healthy pile of cash instead. Selling manure by the tractor bucket is a quick and easy way to make a little extra, but if you take the time to “value add” to your alpaca droppings, you can increase your profits dramatically. Paradoxically, sometimes the smaller the quantity of beans, the bigger the price tag. Chris Reachard of C&C Alpaca Factory in Seaford, Delaware is an expert on ways to profit from poop. He leads hands-on workshops on a regular basis where other farmers can learn from his experience. Here are some of the creative ways he’s found to turn extra muck into a buck. Workshop attendee running dried alpaca beans through a woodchipper before using them to make multiple products.
Manure isn’t just for outdoor gardens - it’s perfect for houseplants, too. Chris packages dried, ground beans as a “magic dust” sprinkle for indoor plants. Compost “tea”, both in concentrate and diluted form, is also perfect for indoor and outdoor use as a foliar feed. The rustic burlap tea bags are a nice touch.
Do be creative.
Cute and whimsical sells, even if it’s poop. Be sure to feature your “bean machines” on your labels.
Do be sure to note the benefits of using alpaca manure in your
garden. Aged manure adds tilth to the soil, making it a better home for both micro- and macrobiota.
Do note that alpaca manure is a good source of carbon, nitrogen, and some trace minerals, although in lower amounts than a commercial fertilizer.
Do let your customers know that raw alpaca manure will not burn their plants - but should be handled with care and applied well before harvest to avoid any potential issues with pathogens.
Don’t be afraid to think outside the box a little! One of the products Chris makes is fire logs. Yes, dried poop is molded into a log for burning (don’t worry, they’re odorless). What could be more eco-friendly than that? You can find plastic brick molds on eBay. If you’d like to start smaller, why not try molding seed starter blocks? Chris makes his with alpaca beans mixed with coco coir. What a great, local alternative to the cow manure versions being sold in stores. Find the mold at http://www.johnnyseeds. com/p-8088-hand-held-20-soil-blocker.aspx.
Don’t make promises you can’t keep.
Talk about the benefits of your product, but don’t make guarantees or unrealistic claims.
Don’t call your manure fertilizer or compost.
Both have specific legal definitions that vary by state. Call it aged manure or just paca poop.
Don’t forget to label your compost “tea” as “Not intended for human consumption.” Better safe than sorry.
Don’t forget to include appropriate disclaimers, for example:
“This manure may contain foreign objects. Please wear gloves when handling all organic material.” Photographs courtesy of Chris Reachard
The easiest way to sell your alpaca poop - by the tractor bucket! If you can’t load, let people come and shovel their own. Or have prefilled amounts available. Five gallon buckets or repurposed feed sacks work very well for most people’s needs.
Craigslist is free and widely seen. Or even a little sign at the end of your driveway will bring customers to you.
Don’t be afraid to barter.
Photographs courtesy of Bag End Suri Alpacas
A budding CSA is trading me summer vegetables for all of the manure they hauled away this spring.
Don’t give your manure away. are very willing to pay for it.
It’s a valuable resource and people
Don’t call your manure organic.
It almost certainly isn’t.
Beans, beans, the magical poop, the more you use, the better your fruit... Selling manure sounds about as easy as can be, but there are some things to take into consideration before you begin cashing in on your beans. Be very careful about what claims you make. Organic, fertilizer, and compost all have specific, legal definitions that vary from state to state. No one is policing the sale of aged alpaca manure, but there are legal repercussions for selling fertilizer that does not list lab-tested NPK values, for example. Similarly, state agencies regulate the sale of compost, as well as the use of the term “organic.” Few, if any, alpaca farms will qualify as organic, particularly those in meningeal worm territory who perform monthly ivermectin prophylaxis. In addition to not using chemical dewormers, farms must use all organic hays and feeds to comply with label requirements. Talk to your hay supplier(s) before you begin selling your manure. Some herbicides used on hay fields are persistent, and manure from animals fed these hays will kill or injure garden plants, making you liable for damages. These herbicides can remain active in treated hay for as long as three years when the hay is stored properly. Similarly, breakdown may take months to years in manure and compost piles. These persistent herbicides contain aminopyralid, clopyralid, or picloram (pyridine carboxylic acids). Current common trade names may include: Curtail, Forefront, GrazonNext, Grazon, Milestone, Redeem and Surmount. New products appear regularly, so always check the ingredients list for pyridine carbolic acids. Be sure you know what is being used on the hays you feed your animal. For more information, read the NC State University Agricultural Extension publication “HERBICIDE CARRYOVER IN HAY, MANURE, COMPOST, AND GRASS CLIPPINGS: Caution to Hay Producers, Livestock Owners, Farmers, and Home Gardeners” available online at http://content.ces. ncsu.edu/herbicide-carryover.
Viral Encephalitis Diseases in Camelids:
Eastern Equine Encephalitis, West Nile Virus, and Equine Herpes Virus by Dr. Daniela Bedenice, DVM South American camelids have been found to be vulnerable to a number of encephalitis diseases found in other livestock. Encephalitis refers to the acute swelling of the brain that is characteristic of this group of diseases, all of which are caused by viruses. As discussed below, vaccination has produced mixed results in clinical trials. Good supportive care is critical for the recovery of animals affected by any of these viral conditions, but the prognosis is typically very guarded and loss is common. Wherever possible, owners should consider taking steps to reduce the risk of exposure to these conditions, as discussed in the accompany sidebar.
Eastern Equine Encephalitis (EEE)
Colorized transmission electron micrograph of the salivary gland of a mosquito infected with the EEE virus (shown in red).
Fred Murphy and Sylvia Whitfield - CDC Public Health Image Library Image #7057, Public Domain, https://commons.wikimedia.org/w/index.php?curid=901852
Eastern Equine Encephalitis (EEE) is a highly fatal alpha-viral disease (family Togaviridae, genus Alphavirus), that has been leading to low numbers of documented fatalities in camelids since at least 2005. The EEE virus is transmitted by the bite of an infected mosquito vector. It is not transmitted directly from animal to animal. A case series in 2007 identified EEE in eight 8 alpacas and one llama in the eastern United States (Nolen-Walston 2007). Interestingly, most of these cases were in young animals (mean age: 2.27 years), with seven of nine patients under one year of age and four of nine animals less than 10 weeks old.
Clinical Signs Clinical signs of EEE infection in the llamas and alpacas were consistent with intracranial disease and included depression, fever, ataxia, seizures, recumbency, vestibular signs and opisthotonus (Nolen-Walston 2007). Despite intensive medical therapy, the treatment of EEE in clinically ill animals is generally unrewarding, with an estimated fatality rate of 90%. (Bedenice 2009; Whitehead 2009). EEE symptoms are similar to those of other viral encephalitis diseases, and overlap with those of thiamine deficiency, listeriosis, and many others. Veterinary involvement in any case where neurological symptoms are present is highly recommended. In October 2005, the prevalence of naturally occurring EEE antibodies was estimated to be 10% in New Hampshire camelids (7/73; geometric mean titer (GMT): 1 – 2.6), based on a serum survey of 73 clinically healthy unvaccinated alpacas. Previous studies had identified a similar prevalence of antibodies to EEE virus in free ranging wildlife. Although the use of serologic findings to establish information on distribution of viral diseases has substantial limitations, seroprevalence surveys may still be useful in detecting geographic areas where the disease is considered to be active. Vaccination Efficacy In 2007, a clinical study published the serum antibody response in two groups of healthy alpacas (n=39 and n=86, respectively) following vaccination with a bivalent killed Equine Encephalitis vaccine (Encevac, Intervet Inc). The EEE vaccine was administered using three intramuscular doses (1ml each) at 4-week intervals (Bedenice 2009; Whitehead 2009). Peak vaccine titers were observed two weeks following the second booster vaccination (i.e. week 10). At this time 76% (study group I) and 88 % (study group II) of all alpacas were seropositive for EEE, based on a plaque reduction neutralization assay (National Veterinary Services Laboratories [NSVL], 1800 Dayton Ave, Ames, IA 50010). Median geometric mean titers (GMT) were not significantly different between huacaya and suri or between male and female alpacas. However, the GMT and the percentage of seropositive animals were significantly higher in juvenile animals (< 2 years old, median GMT: 1.67 – 2.1) than in animals > 6 years old (median GMT: 0.2 – 1.29). Juvenile alpacas thus achieved EEE antibody titers comparable to a report in horses (Holmes 2006), although two booster vaccinations were required in camelids. The latter study did not assess whether neutralizing antibody responses in camelids are associated with vaccine efficacy. However, vaccination with a killed equine vaccine is considered protective against EEE in several species, including horses (Waldridge 2003), emus (Tengelsen 2001) and whooping cranes (Olsen 1997).
West Nile Virus West Nile Virus (WNV) is an arthropod-borne virus (arbovirus) most commonly spread by infected mosquitoes. WNV reached its highest incidence in US horses in 2002, with over 15,000 cases reported that year (USDA 2003, Smith 2008). In 2013, a total of 395 cases of WNV infection were reported in veterinary species with the highest incidence in Texas (n=69), Oklahoma (n=41), and Montana (n=32). Clinical Signs Reported clinical signs of WNV disease in alpacas include an acute onset of depression, anorexia, fever, recumbency and opisthotonus, as well as head tremors, asymmetric ataxia, decreased conscious proprioception and altered mental state (agitation and hyperesthesia) (Kutzler 2004a, Dunkel 2004). Cerebral spinal fluid analysis in three affected animals demonstrated an increased protein concentration (95 – 151 mg/dL, reference 31 – 67 mg/dL) and elevated nucleated cell count (12 – 22 cells/μl, reference 0 – 3 cells/ μl) with a predominance of macrophages and small lymphocytes (Dunkel 2004). Although camelids are considered to be at low risk of developing clinical signs of WNV infection, the observed fatality in affected animals displaying neurological signs is high (Kutzler 2004b, Dunkel 2004). Vaccination Efficacy Preventative encephalitis vaccines are currently not licensed for use in camelids. However, the serological response to an intramuscular formalin-inactivated equine WNV vaccine (West Nile Innovator, Fort Dodge Animal Health, Fort Dodge, Iowa) has been evaluated in 28 alpacas and 56 llamas (Kutzler 2004b). All alpacas and llamas received three vaccinations at 3-week intervals, with fifty-five llamas being vaccinated four times, three weeks apart. No local or systemic adverse reactions were observed in these animals, although adverse reactions, ranging from minor modular swellings at the injection site to anaphylaxis have been previously documented (Kutzler 2004b). Twenty-seven of 28 (96%) alpacas and 26 of 29 (90%) llamas were seropositive three weeks after administration of the second vaccination, with all 28 alpacas and 97% llamas being seropositive three weeks after the third vaccine. Mean GMTs ± SD of seropositive animals following the second vaccination were 169 ± 4 for alpacas and 97 ± 5 for llamas. Following the third vaccination, mean titers increased to 545 ± 4 and 446 ± 4, respectively. This study further reports that the humoral response to WNV vaccination in alpacas persisted for > 40 weeks. The authors concluded that administration of three vaccinations to camelids generally resulted in virus-neutralizing antibody titers similar to those observed after two vaccinations in horses. To date, the efficacy of WNV vaccination of camelids has not been evaluated. However, research in horses has shown that 100% (8/8) of horses vaccinated twice with the same inactivated WNV vaccine used in the camelid study survived and were protected from the onset of WNV encephalitis and viremia. All unvaccinated control horses died, showing severe neurological signs, viremia, and moderate to severe histopathologic lesions in the brain and spinal
cord. Nonetheless, mild to moderate neurological deficits (grade 1 or 2 and less) were also observed for two consecutive days in a few of the vaccinated horses (Seino 2007). It has been suggested that protection of clinical WNV disease may be similarly achieved in some vaccinated camelids (Kutzler 2004b).
Equine Herpes Virus (EHV-1)
In 1984, infection with a herpesvirus indistinguishable from EHV-1 was associated with irreversible blindness in 21 alpacas and one llama of a herd of approximately 100 camelids (Rebhun 1988). Most affected animals had evidence of vitritis, retinitis and optic neuritis in the acute phase of disease. Ophthalmologic findings ranged from no detectable ocular abnormalities to severe hemorrhagic vitritis and chorioretinitis with retinal detachment. A few affected alpacas experienced retrobulbar optic neuritis, causing blindness and midriasis without ophthalmoscopic lesions. Four of the blind animals also developed signs of neurological dysfunction, including depression, nystagmus, head-tilt and /or paralysis. Clinical Signs In a subsequent study, three llamas were intranasally infected with EHV-1, isolated from the brain of an alpaca that had experienced severe neurologic disease (House 1991). Two of the three llamas developed severe neurological disorders 6-8 days following EHV1 inoculation, including head pressing, opisthotonos, blindness, crossed or splayed forelimbs, staggering, ear fasciculations, head tremors and depression. The third llama only demonstrated mild neurological signs during the course of infection. Histopathological lesions in the brain included severe multifocal acute vasculitis with acute cerebral and retinal edema and a moderate multifocal acute meningoencephalitis. Ocular lesions were only observed in one animal, manifesting as moderate acute multifocal necrotizing retinitis with intra-nuclear inclusion bodies. Based on these observations, EHV-1 is believed to gain access to the brain via the olfactory nerve in camelids, while retinal invasion may occur subsequent to olfactory tract infection and/or early viremia (House 1991). In horses, EHV-1 spreads via exposure to nasal or cough secretions, or exposure to fetuses aborted due to the virus, or contaminated fetal fluids or placentae. Similar modes of transmission are suspected in camelids. Vaccination Efficacy Even though EHV-1 infection may result in severe clinical signs in camelids, a low disease prevalence is expected as long as exposure to equids is limited. A serological survey of 270 llamas conducted in Oregon, only detected antibodies to EHV-1 in one animal (Mattson 1994). Routine vaccination against EHV-1 is therefore not recommended. Although unpublished reports have suggested that vaccination with a killed-virus EHV-1 product (Pneumobort-K, Fort Dodge, IA) may induce antibodies in llamas when used according to the manufacturer’s instructions, this vaccination procedure requires further investigation. Modified live-virus EHV-1 vaccines are not recommended in SAC (Mattson 1994). Equine herpesvirus-1 infection is ubiquitous in most horse populations, with the neurotrophic EHV-1 variant showing an increased neuropathogenic potential. Unfortunately, there are only
few studies to date which help determine the utility of current vaccines in preventing naturally occurring cases of Equine Herpes Myeloencephalopathy (EHM). Overall, available killed EHV-1 vaccines may induce variable virus neutralizing antibody responses, depending on vaccine type. Vaccines licensed only for protection against respiratory disease usually have lower antigen content and generate lower serum neutralizing antibody responses. However, both Pneumabort-K® (Pfizer Animal Health) and Prodigy™ (Intervet Schering-Plough Animal Health), have as much as five times the antigen content of their respiratory vaccine counterparts, and both vaccines will thus generate much higher antibody responses to vaccination. In the experimental setting, EHV shedding from the nasopharynx is often significantly reduced in horses having received a modifiedlive virus (MLV) vaccine. While vaccination also increases serum antibody titers, this often fails to alter the duration of cell-associated viremia. However, the control of cell-associated viremia is important for the prevention of EHV-1 associated neurological disease. A recent study (Goehring 2010) evaluated the control of EHV-1 viremia and nasal shedding by commercial vaccines in horses. Three groups of 8 yearling ponies either received 3 treatments with a MLV vaccine (Rhinomune, Boehringer Ingelheim Vetmedica), a killed vaccine (Pneumabort-K, Pfizer Animal Health) or a placebo (control group) followed by viral challenge. Both vaccines offered some level of protection by reducing clinical signs of respiratory disease and decreasing nasal viral shedding. The killed vaccine also suppressed days of viremia. However, neither vaccine completely prevented viremia, and thus the risk of EHM or abortion that can follow. The expectation of vaccination should be to assist with disease control, to reduce severity of clinical signs and to limit virus shedding from infected horses, rather than producing immunity. Selected references:
1. Nolen-Walston R, Bedenice D, S, Rodriguez C et al. Eastern equine encephalitis in 9 South American camelids. JACVIM. JulAug;21(4):846-52, 2007 2. Bedenice D, Bright A, Pedersen D, Dibb J. Humoral response to Equine Eastern Encephalitis vaccination in healthy alpacas. JAVMA 2009;234(4):530-4. 3. Whitehead CE, Bedenice D. Neurologic diseases in llamas and alpacas. Vet Clin North Am Food Anim Pract. 2009;25(2):385-405 4. Tengelsen L, Bowen R, Royals M, et al. Response to and efficacy of vaccination against eastern equine encephalomyelitis virus in emus. J Am Vet Med Assoc 2001;218:1469-1473. 5. Waldridge B, Wenzel J, Ellis A, et al. Serologic responses to eastern and western equine encephalomyelitis vaccination in previously vaccinated horses. Vet Ther 2003;4:242-248. 6. Holmes MA, Townsend HG, Kohler AK, et al. Immune responses to commercial equine vaccines against equine herpesvirus-1, equine influenza virus, eastern equine encephalomyelitis, and tetanus. Vet Immunol Immunopathol 2006;111:67-80.
EEE virus in domestic and feral swine in Georgia. J Vet Diagn Invest 1996;8:481-484. 10. Hoff GL, Issel CJ, Trainer DO, et al. Arbovirus serology in North Dakota mule and white-tailed deer. J Wildl Dis 1973;9:291-295. 11. Yaeger M, Yoon KJ, Schwartz K, Berkland L. West Nile virus meningoencephalitis in a Suri alpaca and Suffolk ewe. J Vet Diagn Invest. 2004;16(1):64-6. Kutzler MA, Bildfell RJ, Gardner-Graff KK, Baker RJ, Delay JP, Mattson DE. West Nile virus infection in two alpacas. J Am Vet Med Assoc. 2004;225(6):921-4, 880. 12. Dunkel B, Del Piero F, Wotman KL, Johns IC, Beech J, Wilkins PA. Encephalomyelitis from West Nile flavivirus in 3 alpacas. J Vet Intern Med. 2004;18(3):365-7. 13. Kutzler MA, Baker RJ, Mattson DE. Humoral response to West Nile virus vaccination in alpacas and llamas. J Am Vet Med Assoc. 2004;225(3):414-6. 14. Seino KK, Long MT, Gibbs EPJ et al. Comparative Efficacies of Three Commercially Available Vaccines against West Nile Virus (WNV) in a Short-Duration Challenge Trial Involving an Equine WNV Encephalitis Model. Clin Vaccine Immunol. 2007; 14(11): 1465–1471. 15. Sellon DC, Long MT. Equine Infectious Diseases. Saunders Elsevier; 1st ed; 2007. 16. Rebhun WC, Jenkins DH, Riis RC, Dill SG, Dubovi EJ, Torres A. An epizootic of blindness and encephalitis associated with a herpesvirus indistinguishable from equine herpesvirus I in a herd of alpacas and llamas. J Am Vet Med Assoc. 1988;192(7):953-6. 17. House JA, Gregg DA, Lubroth J, Dubovi EJ, Torres A. Experimental equine herpesvirus-1 infection in llamas (Lama glama). J Vet Diagn Invest. 1991;3(2):137-43. 18. Mattson DE. Update on llama medicine. Viral diseases. Vet Clin North Am Food Anim Pract. 1994 Jul;10(2):345-51. 19. Goehring LS, Wagner B, Bigbie R et al. Control of EHV-1 viremia and nasal shedding by commercial vaccines. Vaccine. 2010 Jul 19;28(32):520311.
Dr. Daniela Bedenice is a leading expert on llamas, alpacas and other camelid species at the Cummings Veterinary School at Tufts University. Dr. Bedenice grew up in rural Germany, where she trained and cared for many horses. After earning her veterinary degree from the Free University of Berlin, Dr. Bedenice undertook a residency with a private practice. Soon after, she moved to the United States, earned her board certification in both veterinary internal medicine and emergency and critical care, and joined the faculty at the Cummings School. As an Assistant Professor in the Department of Clinical Sciences at the Cummings School of Veterinary Medicine,Dr. Bedenice leads and contributes to a variety of courses and clinics, including large animal internal medicine, gastro-intestinal disease and neurology, camelid medicine, gastro-intestinal pathophysiology, neuro-pathophysiology, toxicology, clinical pharmacology. She also serves as a facilitator for the Problem-Based Learning course.
7. Olsen GH, Turell MJ, Pagac BB. Efficacy of eastern equine encephalitis immunization in whooping cranes. J Wildl Dis 1997;33:312-315. 8. Tate CM, Howerth EW, Stallknecht DE, et al. Eastern equine encephalitis in a free-ranging white-tailed deer (Odocoileus virginianus). J Wildl Dis 2005;41:241-245. 9. Elvinger F, Baldwin CA, Liggett AD, et al. Prevalence of exposure to
Know the Enemy -
Common Mosquito Vectors and How to Control Them Mosquitos are everywhere. There are over 2,500 species of mosquito worldwide, and at least 150 species in the United States. Most carry diseases with the potential to infect humans or their livestock. In North America, the West Nile virus is carried by several species of mosquito in the Culex genus, while Eastern Equine Encephalitis is primarily found in Culiseta melanura. Both of these diseases have the potential to infect alpacas. PRIMARY VECTOR SPECIES Culex mosquitos, like most species, lay their eggs in water. Culex nigripalpus is the predominant pest species in the southeastern United States into the southern half of Texas. Females prefer to lay their rafts of 90-120 eggs in recently flooded areas like roadside ditches and agricultural fields, where the eggs hatch in 24-36 hours. Larval development to the production of an adult takes seven to 14 days depending on water temperature. Newly hatched C. nigripalpus females begin seeking blood meals within 48 hours. In dry conditions, this species primarily feeds on birds in damper, forested areas; during wetter periods, the mosquitos forage more widely and seek out diverse mammalian species. In the northeast and midwest, Culex pipiens is one of the most important mosquito vectors. Females lay rafts of 200-300 eggs in stagnant water as often as every third day; development from egg to pupa takes one to two weeks, with pupation lasting from four days to several weeks, depending on conditions. Females that hatch late in the summer will seek shelter (often in residences, hence the species common name “house mosquito”), and enter a state of suspended metabolism until spring, when they will again seek water in which to lay eggs. Both Culux pipiens and C. tarsalis are important vectors for WNV transmission on the West Coast. C. tarsalis in particular feeds more often on mammalian Culex species larvae. sources than C. pipiens. It is considered a “rural” mosquito, preferring to By (Image: James Gathany, CDC) - A New Model for Predicting Outbreaks of West Nile Virus. Gross L, PLoS Biology Vol. 4/4/2006, e101. http://dx.doi.org/10.1371/journal.pbio.0040101. lay eggs in watered fields, rice paddies, and similar vegetated areas. Larval See also: Kilpatrick AM, Kramer LD, Jones MJ, Marra PP, Daszak P (2006) West Nile Virus development to adult can take as little as eight days up to several weeks, with Epidemics in North America Are Driven by Shifts in Mosquito Feeding Behavior. PLoS Biol 4(4): e82 doi:10.1371/journal.pbio.0040082, CC BY 2.5, https://commons.wikimedia.org/w/ faster development at higher temperatures. However, survivability is lowered index.php?curid=1441809 below 70°F and above 95°F. This species tolerates low levels of salinity. They are strong fliers, and can travel 10 miles or more from where they hatch. Like C. pipiens, C. tarsalis can overwinter in sheltered buildings, caves, and other protected locations. Culex tarsalis is also the primary vector species for Western Equine Encephalitis (WEE) transmission on the West Coast. WEE is closely related to EEE, and causes similar symptoms. Culiseta melanura, the black-tailed mosquito, is the primary vector species for EEE. This mosquito is endemic from Quebec to Florida, and as far west as Texas. It can overwinter in its larval form, unlike most mosquitos that overwinter as eggs or adults. Rafts of 100+ eggs are laid in sheltered bodies of water (hollow stumps, at the base of flooded trees, etc.) multiple times per season in its southern range, while northern populations may only lay twice. If the larvae hatch into cold weather, they hibernate in bottom sediments and complete development the following year. CONTROL MEASURES Large scale mosquito control measures are often undertaken by state and local municipal governments. They may include the use of “adulticides,” insecticides that target adult mosquitos; “larvicides,” insecticides that kill immature stages; water surface films, that disrupt water surface tension and suffocate larvae; or biological control agents that target either or both. Timing and location of insecticide use are dictated by the biology of the species being targeted, weather conditions, and local land use. There are potentially adverse effects to widespread insecticide application - even biological controls such as Bacillus thuringiensis (Bt) affect non-target species, and all adulticides are broad spectrum in nature. Many insecticides are toxic to aquatic insects, including dragonfly and damselfly nymphs; other aquatic organisms, particularly fish; beneficial insects, including honeybees; and may be toxic to birds and mammals, although more study is required. Clearly, there are potential drawbacks to conventional control measures. As a farm owner, there are many mosquito control measures that can be taken short of using chemical insecticides. First and most importantly, remove standing water sources - even tiny ones! As noted above, mosquitos can complete the transformation from egg to
adult in a very short time period, and only require small amounts of water. One critical step is improving field drainage to avoid standing water, as well as mowing long patches of grass and weeds that trap moisture (this helps eliminate other biting flies, too). Be sure to remove containers that hold even small amounts of water - empty those buckets, get rid of old tires, and store your wheelbarrow upside down. Regularly emptying water troughs and bird baths and replenishing them will all reduce the number of breeding mosquitos in your immediate area. If troughs or other containers cannot be emptied, the addition of goldfish, mosquitofish, or Bt will reduce or eliminate larval mosquitos. Although bats and purple martins sadly do not eat as many mosquitos as has been advertised, they do consume some as part of their diet, and there is a benefit to encouraging them and other natural mosquito predators. These include dragonflies and damselflies, which are voracious predators on mosquitos in both their larval and adult forms. Small fish also eat mosquito larvae eagerly. A healthy pond with good water movement promotes an ecosystem where mosquitos are naturally held in check - stagnant water without inhabitants is prime breeding ground for mosquitos and nothing else. For adult mosquito control, bug zappers have been shown to be absolutely ineffective - and studies show that they may zap more Aedes mosquito on human host. good bugs than bad. The jury is out on machines that claim to By (Image: James Gathany, CDC) - A New Model for Predicting Outbreaks of West Nile Virus. Gross L, PLoS attract and kill mosquitos only, usually by emitting heat and CO2. Biology Vol. 4/4/2006, e101. http://dx.doi.org/10.1371/journal.pbio.0040101. See also: Kilpatrick AM, Kramer LD, Jones MJ, Marra PP, Daszak P (2006) West Nile Virus Epidemics in North America Are Driven by Shifts No empirical studies exist that verify manufacturer’s claims. While in Mosquito Feeding Behavior. PLoS Biol 4(4): e82 doi:10.1371/journal.pbio.0040082, CC BY 2.5, https:// these machines may be useful for clearing a limited area for a limited commons.wikimedia.org/w/index.php?curid=1441809 time, it is unlikely that they make a significant dent on anything more that a purchaser’s wallet. Personal ultrasonic devices that are supposed to frighten mosquitos by mimicking the hum of a dragonfly’s wings fail for the simple reason that mosquitos do not avoid dragonflies. Humans should carefully consider the merits of various insect repellents and use them during peak times of mosquito activity. While large scale application of mosquito repellents is likely impractical for most farms, strategically locating fans throughout alpaca resting areas can create safe havens for alpacas when they are not grazing. Mosquitos are most active at dusk and dawn - if your area is at high risk for mosquito borne diseases, keeping your alpacas in during these times will help minimize their exposure. Using yellow lights in the barn will also attract fewer insects inside than white lights. REFERENCES Laura B. Goddard, Amy E. Roth, William K. Reisen, and Thomas W. Scott, “Vector Competence of California Mosquitoes for West Nile Virus,” Emerging Infectious Diseases 2002 Dec; 8(12) Goudarz Molaei, Theodore G. Andreadis, Philip M. Armstrong, John F. Anderson, and Charles R. Vossbrinck, “Host Feeding Patterns of Culex Mosquitoes and West Nile Virus Transmission, Northeastern United States,” Emerging Infectious Diseases 2006 Mar; 12(3): 468–474 Edward B. Hayes, Nicholas Komar, Roger S. Nasci, Susan P. Montgomery, Daniel R. O’Leary, and Grant L. Campbell, “Epidemiology and Transmission Dynamics of West Nile Virus Disease,” Emerging Infectious Diseases 2005 Aug; 11(8): 1167 T.C. Thiemann, D.A. Lemenager, S. Kluh, B.D. Carroll, H.D. Lothrop and W.K. Reisen, “Spatial Variation in Host Feeding Patterns of Culex tarsalis and the Culex pipiens complex (Diptera: Culicidae) in California,” J Med Entomol. 2012 Jul; 49(4): 903–916. http://www.metapathogen.com/mosquito/culex/ http://www.ipm.ucdavis.edu/PMG/PESTNOTES/mosquitofarm12.html?printpage http://entnemdept.ufl.edu/creatures/aquatic/Culiseta_melanura.htm http://emedicine.medscape.com/article/233568-overview http://www.fws.gov/cno/refuges/donedwards/CCP-PDFs/AppendixK4_EffectsofMosquitoControl.pdf http://mosquito.ifas.ufl.edu
Haemonchus contortus, known to many as barber pole worm, is one of the leading causes of death in small ruminants world wide. Following are two articles addressing the use of fecal egg counts and FAMACHA anemia scoring as means for reducing the impact of this parasite on your herd. Although these articles were not written specifically with alpacas in mind, their contents apply to South American camelids. If you were not one of those fortunate enough to hear Bob speak at the 2015 Suri Summer Symposium, here are some of the resources you need to combat internal parasites in general and Haemonchus in particular. Â
FECAL EGG COUNTS: USES AND LIMITATIONS Bob Storey, MS, RVT Assistant Research Scientist Department of Infectious Disease University of Georgia College of Veterinary Medicine Athens, GA USA
Abstract: The microscopic examination of feces for the assessment of gastrointestinal parasite infection has been a mainstay of clinical and research parasitology labs for many decades. Even with the widespread use of fecal egg counts (FECs) in the medical and scientific community, the routine use of FECs by farmers and producers is quite limited in most areas of the world. The lack of use of this very valuable tool is most probably due to a lack of consistent and understandable information regarding the simplicity and value of the FEC. Some of the information that one can obtain via the internet concerning FECs is of variable quality and clarity. Accurate information and training can help to correct the misunderstandings concerning the limitations of the FEC as well as expand its acceptance and use. The modified McMaster FEC technique, which is one of the most widely used quantitative FEC methods in practice today, simple to perform and when used as an adjunct to body condition score, FAMACHA score, geographical location, and fecal consistency scoring (e.g. The Five Point Check) can provide a wealth of information to the farmer/producer. The quantitative FEC not only provides the trained user with information regarding the types parasites present in the sample (trichostrongyles, tapeworm, whipworm, coccidia, lungworm, etc.) as well as an estimate of the quantity of parasite eggs being shed in the feces (eggs per gram) for monitoring pasture contamination. The FEC is also invaluable in the monitoring of anthelmintic (drench/dewormer) effectiveness in controlling these parasites in the flock/herd. The McMaster FEC, when performed with reasonable care and consistency, provides the user invaluable information pertaining to parasite control and management, which can lead to improved herd health and increased production.
Introduction and historical perspective: The microscopic examination of feces for the detection of gastrointestinal parasite eggs as an indicator of parasite infection is one the most widely used tools in classical clinical parasitology as well as parasitology research labs. The FEC has also gained popularity as a valuable tool among producers and producer groups. A search of the published scientific literature shows that there have been more than 6000 scientific papers (all species and disciplines) published using data from fecal egg counts since the appearance of the first publication on the subject in 1923. The 1923 article was entitled “Investigations on the control of hookworm disease. XV. An effective method of counting hookworm eggs in feces” and was authored by Dr. Norman Stoll (Stoll, 1923) . Stoll developed a quantitative method for hookworm eggs (in humans) while working at the School of Hygiene and Health at John’s Hopkins University. The procedure, "Stoll dilution egg-counting technique", created by Stoll was adopted around the world for major epidemiological studies of hookworms. This significant contribution also provided the genesis from which current fecal egg count procedures and techniques evolved (Ashton, 1977). Stoll would follow his human hookworm work with additional publications, but his publication in 1930, “On Methods of Counting Nematode Ova in Sheep Dung” helped to launch the quantitative fecal egg count into the arena of veterinary medicine (Stoll, 1930). Many others have built upon Stoll’s technique over the years, with one of the most significant occurring in 1939, where H.V. Whitlock was serving as a laboratory assistant for the McMaster Animal Health Laboratory in Sydney Australia. Whitlock, in the course of his duties, performed hundreds of fecal egg counts every day, using the method described by Stoll in his 1930 paper and sought a way to improve his lab efficiency. Whitlock developed a special slide that incorporated Stoll’s precise sampling with a flotation technique (Whitlock and Gordon, 1939). The resulting “McMaster Counting Chamber” along with the modifications Whitlock would later make (Whitlock, 1948), are the basis for the many Modified McMaster fecal egg count slides and procedural variants widely used today. What is a Fecal Egg Count? History aside, what exactly are fecal egg counts? (Note-depending on the country/locale that you are in, a fecal egg count may be referred to as a “fecal”, “FEC”, “epg”, “worm egg count”, “WEC”, “ fecal worm egg count”, “worm test” or just “egg count” – ergo “A rose by any other name would smell as sweet”?). A FEC is a procedure performed on a manure sample to detect the presence of parasitic worm eggs. There are two classes of FEC, one being qualitative, meaning that the results are reported as “positive” or “negative” and are generally based on a basic
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fecal floatation procedure. Qualitative FECs can also be reported with a minus sign ( - ) for negative (no eggs seen) or positive as “+, ++, +++”, with the number of plus signs signifying the subjective opinion of the technician as to the number of eggs present. The qualitative FEC is generally performed by mixing a small amount of feces with a floatation solution in a small vial. The solution level is increased to the point where a small positive meniscus is formed and then a microscope slide cover slip is then placed on top. This is allowed to sit for 10 to 30 minutes depending on the protocol that is being followed, after which time the cover slip is carefully lifted from the vial and placed on a microscope slide for examination with a microscope The entire coverslip is examined for the presence of parasitic worm eggs under a magnification of 100x (typically)(Hansen and Perry, 1994) (Figure 1). .
Figure 1. Fecal Float. A) Feces and floatation solution mixture in vial with positive meniscus. B) Cover slip carefully placed on vial. C) Allow to sit for 10 ‐ 30 minutes. D) Carefully remove coverslip. F) Place on slide. G) Positive float or +. H) Positive float moderate or ++. I) Positive heavy or ++++.
The second class of FEC is the quantitative FEC. Quantitative FEC results are reported in eggs per gram (epg) of manure. The most common method of quantitative FEC for sheep and goats is the Modified McMaster technique mentioned in the introduction section. Although there are several variations of the Modified McMaster procedure (Coles et al., 1992; Cringoli et al., 2004; Foreyt, 2001; Ministry of Agriculture, 1977; Zajac and Conboy, 2012), all of the various methods use a weighed fecal sample, a known volume of flotation solution, and the specialized McMaster counting slide (Figure 2A). The two chambers of the slide are filled with the manure/flotation mixture and then the trichostrongyle type eggs under the two McMaster chamber grids are counted (Figure 2B).
Figure 2 A) McMaster slide being filled with flotation mixture. B) View through a compound microscope at 100X of McMaster chamber with trichostrongyle eggs present. Non trichostrongyle eggs, such as tapeworm, whipworm, and coccidian oocysts are noted, but not actually counted (See Figure 3). The total trichostrongyle type eggs counted under both grids are multiplied by a dilution factor that is determined by the concentration of feces in the floatation solution. This dilution factor is procedure/protocol specific and is determined by the weight of the feces, the volume of the floatation solution that the feces were dissolved in, and the volume of this mixture visible under both of the McMaster chamber grids. This may sound a little complicated, but quantitative McMaster counts are no more difficult to perform than simple flotations, and the equipment is relatively inexpensive and reusable – many producer cooperatives and breed groups that I work with purchase a microscope and McMaster slides for the group, and allow the members to share them. An equipment list and detailed instructions for performing fecal egg counts can be found on the American Consortium for Small Ruminant Parasite Control (ACSRPC) web site at www.wormx.info or www.acsrpc.org.
Figure 3 Common eggs and oocysts of small ruminants
What does a FEC tell us? A single FEC provides us with an estimate of the number of target parasite species eggs present in a particular fecal sample. Since our major interest is sheep and goats, our target species are the Trichostrongyles, which includes any worm of the genus Trichostrongylus; however, since we are mainly interested in sheep and goats for our discussion the term trichostrongyles will be limited to Haemonchus spp., Trichostrongylus spp., Ostertagia spp., Teladorsagia spp., Oesophagostomum spp., and Cooperia spp.. It is not possible to visually differentiate the eggs of the aforementioned species accurately due to the similarity in of the size and shape of their eggs, so the epg value determined by FEC for a sheep or goat is only for these trichostrongyles. Other species that can be seen on FEC that can be readily identified (and quantified if desired) are Nematodirus spp., Chabertia spp.,Bunostomum spp, Strongyloieds spp, Trichuris spp. (whipworm), Monezia spp. (tapeworm), Eimeria spp. (coccidia), Marshallagia spp, and Capillaria spp. The species listed in this section is not by any means intended to be complete, but does reflect the most commonly encountered species in sheep and goats (Foreyt, 2001). Another important fact concerning FECs is that just because no eggs are detected on the slide, does not mean the animal is free of gastrointestinal parasites. The failure to see eggs can result not only from there being only a few parasites present in the animal or using an egg counting technique that is not sensitive enough, but also from the random chance no eggs were deposited in the particular portion of feces collected as a sample. Also, one must bear in mind that the egg per gram (epg) value determined by a FEC represents a snapshot in time of the specific fecal sample tested for a given animal, and that it in actuality tells us very little about the actual worm burden of the animal. Logic would suggest that a high egg count means a high worm count, but we are dealing with a biological system where many factors affect the actual egg production rate of female worms. First and foremost, different parasite species exhibit differing rates of fecundity (eggs production) as can be seen in Table 1. Nematode Daily egg production/female 5000-15000 Haemonchus 100 - 200 Ostertagia, Trichostrongylus 1000-3000 Cooperia 50-100 Nematodirus 5000-10000 Oesophagostomum, Chabertia Table 1. Daily egg production ranges of female trichostrongyles Source (Hansen and Perry, 1994). Other factors include the number of mature adult parasites established in the GI tract of the animal, the host animals level of immunity, the age of the host animal, the sex and the pregnancy status of the host animal, the developmental
stage of the parasite infection, the species of parasite(s) present in the host, as well as the consistency of the feces (Düwel, 1990; Hansen and Perry, 1994). Also, in addition to all of the previously mentioned factors affecting parasite egg production, one must also be mindful of the many factors affecting egg distribution in the feces. Digested material does not flow through the alimentary canal at a constant rate, female worms do not lay eggs continuously, nor are they are timed or synchronized with other females in their release of eggs. So, the actual egg count achieved for a given fecal sample depends on a lot of variables. Regardless of all of the above, Haemonchus contortus and Trichostrongylus colubriformis worm burdens are generally considered to correlate with fecal egg count (Cabaret et al., 1998); however, the diagnostic significance of FECs and/or worm burden profiles for the purpose of treatment decisions should not be considered in a vacuum, but should be evaluated in relation to the history and management of the flock and be supported by an assessment of the presence or absence of clinical signs and indications as one would detect with the Five Point Check (Bath et al., 2010). Regardless of all of the caveats and limitations, FECs, when used in context with observation and other common integrated parasite management techniques, do provide us with valuable information for the management of parasites in our herd or flock. So why perform FECs? Regardless of all of the apparent limitations and variability associated with FECs, there are three things that FECs help us determine, and they do so quite well. The first and most important is monitoring the efficacy of your anthelmintics – resistance detection. If a group animals has a high fecal egg count and is treated with a particular drench or combination, and then 10 days later a follow up FEC shows a zero or extremely low FEC (less than 5% of the pre-treatment value) for that group, then you can be fairly assured that your drench or combination worked (Coles et al., 1992). I invoke the inimitable words of Dr. Ray M. Kaplan concerning this fact, “DEAD WORMS DO NOT LAY EGGS”. The second use of FECs is that they can provide information for monitoring pasture contamination. Routine FEC surveillance can provide information to a producer as to how fast the parasite contamination is building up on a pasture. This information can then aid in making decisions as to when to move animals off of a pasture to avoid a potentially dangerous parasite situation, as well as to provide valuable knowledge for determining whether a previously used pasture would be suitable for re-use during that same grazing season. For example if a pasture is grazed for a couple of months early in the grazing season and average FECs were high, then it very likely has a high
level of contamination of infective larvae and therefore would not be suitable for grazing lambs or kids in midsummer. And the third is to aid in selection of animals that exhibit resistance to worms, or exhibit resilience in the face of a worm challenge. Resistance to parasites and resilience in the face of a parasite challenge are both heritable traits (Baker, 1999), and aid in the selection of animals exhibiting these traits. An animal with a consistently low FEC and low FAMACHA scores and rarely needs drenching compared to his herd mates exhibits signs of resistance. But in the same herd, and animal with consistently low FAMACHA scores, good body condition scores, and routinely has FECs in the 10,000 range is resilient and also a heavy pasture contaminator. Both of these animals are productive animals, but one is more desirable than the other. Without the information from the FEC, you would not know whether the trait you were seeing is resistance or resilience (personal experience). Conclusion/Summary: FECs are a widely used technique for quantifying parasite eggs in feces and is used for both clinical parasitology and research parasitology. The FEC is also used by farmers and producers to provide information relevant to parasite control in their sheep flocks and goat herds. The FEC is a simple test procedure that can be either qualitative (yes or no) or quantitative (# of eggs per gram) type. Both provide useful information, but for the sheep or goat farmer the quantitative variety provides more useful information in that all grazing animals have parasites, so the answer to the question “Do my animals have parasites?” is a given. FECs provide a snapshot in time of the number of parasite eggs in a given fecal sample, but does not provide accurate estimation of worm burden in the animal beyond the assumption that a high fecal egg count implies a high worm count. A FEC result that is negative or less than the minimal sensitivity of the test performed does not mean that the animal is free from parasites. The FEC provides its primary value to the farmer or producer as a tool for monitoring anthelmintic / dewormer / drench efficacy – resistance detection, the ability to monitor the rate at which eggs are being deposited on the pasture – pasture contamination, and it also provides additional data for detecting the genetic traits of resistant or resilient animals for animal selection/breeding. The FEC, when used properly, is a very fine tool for the producer to add to his parasite management tool box.
Bibliography Ashton, C.C., 1977. Norman R. Stoll: One of the World's Great Parasitologists. The Journal of Parasitology 63, 878902. Baker, R.L., 1999. Genetic resistance to endoparasites in sheep and goats. A review of genetic resistance to gastrointestinal nematode parasites in sheep and goats in the tropics and evidence for resistance in some sheep and goat breeds in sub-humid coastal Kenya. Animal Genetic Resources Information, 13-30. Bath, G.F., Wyk, J.A.v., Malan, F.S., 2010. Targeted selective treatment of sheep using the Five Point Check©. Journal of Commonwealth Veterinary Association 26, 29-32. Cabaret, J., Gasnier, N., Jacquiet, P., 1998. Faecal egg counts are representative of digestive-tract strongyle worm burdens in sheep and goats. Parasite 5, 137-142. Coles, G.C., Bauer, C., Borgsteede, F.H.M., Geerts, S., Klei, T.R., Taylor, M.A., Waller, P.J., 1992. World Association for the Advancement of Veterinary Parasitology (W.A.A.V.P.) methods for the detection of anthelmintic resistance in nematodes of veterinary importance. Veterinary Parasitology 44, 35-44. Cringoli, G., Rinaldi, L., Veneziano, V., Capelli, G., Scala, A., 2004. The influence of flotation solution, sample dilution and the choice of McMaster slide area (volume) on the reliability of the McMaster technique in estimating the faecal egg counts of gastrointestinal strongyles and Dicrocoelium dendriticum in sheep. Veterinary Parasitology 123, 121-131. Düwel, D., 1990. Studies in ruminants on the incidence of eggs of gastrointestinal nematodes in faeces in relation to worm burden. / Untersuchungen bei Wiederkäuern zum Vorkommen von Eiern gastro-intestinaler Nematoden im Kot in Relation zur Wurmbürde. Mitteilungen der Österreichischen Gesellschaft für Tropenmedizin und Parasitologie 12, 69-80. Foreyt, W.J., 2001. Veterinary Parasitology Reference Manual, 5th Edition. Blackwell. Hansen, J., Perry, B., 1994. Helminth Parasites of Ruminants. International Laboratory for Research on Animal Diseases, Nairobi, Kenya. Ministry of Agriculture, F., and Food., 1977. Manual of veterinary parasitological laboratory techniques. Her Majesty's Stationary Office. Stoll, N.R., 1923. Investigations on the control of hookworm disease. XV. An effective method of counting hookworm eggs in feces. . American Journal of Hygiene, 59-70. Stoll, N.R., 1930. On Methods of counting Nematode Ova in Sheep Dung. Parasitology 22, 116-136. Whitlock, H.V., 1948. Some modifications of the McMaster helminth egg-counting technique apparatus. J. Counc. Sci. Ind. Res, 177-180. Whitlock, H.V., Gordon, H., 1939. A New Technique for Counting Nematode Eggs in Sheep Feces. Journal of the Council for Scientific and Industrial Research 12. Zajac, A.M., Conboy, G.A., 2012. Veterinary Clinical Parasitology, 8th Edition. John Wiley & Sons, Inc., Ames, Iowa , USA.
FAMACHA© and DrenchRite®: Tools for Smart Drenching Bob Storey, MS, RVT Assistant Research Scientist Department of Infectious Diseases University of Georgia, College of Veterinary Medicine Athens, GA 30602 Abstract: Gastrointestinal nematodes (GIN), particularly Haemonchus contortus (barber pole worm), are the leading cause of morbidity, mortality, and subsequent economic loss in small ruminant production in the Southeastern United States (SEUS). Control of GI parasites in small ruminants is complicated by the problem of anthelmintic (dewormer) resistance (AR). The fortunate small ruminant producer has one (at most two) anthelmintics that are still effective against GIN in his herd or flock; however, total anthelmintic failure on small ruminant farms is increasing rapidly. “Smart Drenching” is a concept developed and recommended by the American Consortium for Small Ruminant Parasite Control (ACSRPC). “Smart Drenching” allows a producer to manage GIN in their herd or flock while minimizing genetic selection pressure for anthelmintic resistance. Reduced selection pressure will extend the useful life of their still effective anthelmintics. The Smart Drenching approach utilizes parasite biology, host physiology, dynamics of selection for resistance, and the resistance status of worms on the farm to effectively manage parasites. The FAMACHA© system is an integral part of Smart Drenching and provides a simple and effective tool for identifying animals in need of anthelmintic treatment (where Haemonchus contortus is the primary parasite). The DrenchRite® larval development assay is an in vitro assay that provides the producer with the resistance status of drugs in the benzimidazole, levamisole, and ivermectin families of anthelmintics as well as moxidectin. The DrenchRite® assay also includes identification of GIN species present in the submitted sample. The use of these two tools provides the producer the information necessary for determining which animals to treat, and what dewormer will provide effective treatment. I. Introduction: The trichostrongyle parasites Haemonchus contortus (Hc) and Trichostrongylus colubriformis (Tc) commonly infect small ruminants throughout the United States (USDA-NAHMS, 2011). Hc is a virulent, blood-feeding nematode that
damages the abomasum and abomasal mucosa in sheep and goats. Heavy infections cause profound anemia, hypoproteinemia, loss of fecal consistency, and weight loss. Severely affected animals die from exsanguination (blood loss). Periparturient females, kids, and lambs in their first grazing season, or animals debilitated from another disease are most at risk. Mandibular edema (“bottle jaw”) can occur in severe small ruminant cases, but is not always present in haemonchosis. Hc is generally considered a warm climate parasite, found primarily in the SEUS; however, the 2011 NAHMS sheep survey showed Hc present in 21 of 22 non-SEUS states (USDANAHMS, 2011). See Table I. Additionally, cases of severe to fatal haemonchosis are now being regularly reported from northern states, as well as Canada, confirming that Hc is not as geographically restricted as once believed. Tc resides in the small intestine, and is not as pathogenic as its Hc cousin; however, it can still have deleterious effects (including death) to weak or heavily burdened animals. Morbidity is primarily through irritation of the small intestine/small intestine mucosa. Clinical symptoms include weight loss from malabsorption of protein and anorexia, as well as diarrhea. In contrast to Hc, Tc infections are not generally associated with anemia. Tc is considered a cool climate parasite. A. Anthelmintic resistance (AR) is defined as the ability of certain worms in a population to survive drug treatments that are generally effective against the same worm species and stage of infection. AR results from genetic changes that negate a drug’s mode of action against an individual nematode. Genes that confer resistance to an anthelmintic can exist prior to the first use of the drug, but can also arise from spontaneous mutations due to the immense genetic diversity in GIN populations. Changes in the resistance allele frequency in a worm population occurs as a result of the anthelmintic treatment genetically selecting resistant worms – when you treat an animal with an anthelmintic, you are culling the susceptible worms. Over time and repeated treatment, a growing proportion of the worm population will become resistant to that dewormer, ultimately leading to that dewormer no longer being clinically effective. AR in small ruminant GIN resulting from indiscriminate use and over reliance on chemical dewormers, is now a critical problem in sheep and goats in the US (Howell, Burke, et al, 2008) . AR to the benzamidazole class of dewormers, such as fenbendazole, (e.g., Safeguard®, Panacur®, and Valbazen®, etc.), is ubiquitous in the US, with ivermectin and moxidectin resistance rising rapidly
(See Table II). Treatment failures resulting from AR have led to increased morbidity and mortality in small ruminants. Studies of Diagnostic Lab necropsy reports show that GI parasitism is the number one cause of death for goats in the SEUS. Hc results in more goat fatalities than the next three leading causes combined. State
Hc >90% Avg Positive Hc % Hc 15 12 3 41.2 CA 7 6 4 62.4 CO 14 11 4 62.1 ID 29 29 19 87.8 IA 19 19 14 89.8 KS 9 8 2 41.0 KY 14 14 5 60.6 MI 32 31 19 83.8 MN 6 5 3 73.7 MO 14 13 6 68.9 MT 7 6 0 42.4 NM 14 13 6 68.4 NY 11 11 5 83.3 OH 18 15 3 52.9 OR 8 8 4 86.6 PA 19 18 9 78.4 SD 11 11 11 96.9 TX 34 25 1 26.2 UT 10 10 6 86.0 VA 7 7 1 58.3 WA 21 19 12 74.0 WI 20 18 5 53.6 WY 339 309 142 67.2 22 States Table I: Distribution of Hc in non-southeastern states. Hc was found on 91% of farms surveyed in 21 out of 22 states surveyed. Eighteen states had at least 1 farm with 100% Hc. The overall survey showed a worm population of 67.2% Hc. (USDANAHMS, 2011) Dewormer
AR AR prevalence prevalence (2002-2006) (2002-2011) Benzimidazole 98 99.4 Levamisole 54 54 Ivermectin 76 76.7 Moxidectin 24 39.4 MDR* 48 48.9 TAF** 17 23 Table II: Prevalence of anthelmintic resistance (percent) in sheep in goats for the periods of 2002-
2006 (Howell, Burke, et al, 2008), and 2002-2011 (Kaplan lab, 2013). *MDR = Multi-drug resistance. **TAF = Total anthelmintic failure. AR is therefore an inevitable consequence of anthelmintic use, and as such, AR GIN in sheep and goats is now the rule instead of the exception. Extending the life of anthelmintics by decreasing selection pressure for AR is now the most important challenge facing sheep and goat producers. There is currently only one new anthelmintic in the development/approval process, Zolvix® (monapantel). This drug is not currently approved for use in the US. The market availability date for this new drug, in the US, is currently unknown. At present, we only have the drugs that we have. Reducing AR selection pressure as well as dealing with existing AR parasites requires good animal husbandry as well as intelligent strategies that take advantage of parasite biology and host physiology. These strategies, such as “targeted selective treatment” (TST) and effective dewormer use, coupled with “novel” techniques for parasite management are necessary for the small ruminant farmer to survive in the era of anthelmintic resistance. Failure to adopt common sense principles of parasite management will also lead to the rapid development of resistance to any new drugs that come on the market. B. Parasite Distribution: The natural distribution of parasites across a herd or flock is an important fact that can be used to the producers’ advantage. In healthy sheep and goat herds, about one-third of the animals harbor 80 per cent of the gastrointestinal parasites (Barger, 1985). This “over dispersion” of parasites among hosts justifies selective, rather than whole-herd, anthelmintic treatment. The untreated animals in the herd provide minimal pasture contamination, but are a major source of Refugia (worms that are not under selection pressure from exposure to an anthelmintic) as shown in Figure I. Refugia plays an important role in how quickly a given worm population will become resistant to an anthelmintic (Dobson, Hosking, et al, 2012). Without Refugia, only resistant worms are left to reproduce with each other, which rapidly amplify resistant genes in the worm population. Accurate identification of animals that require anthelmintic treatment to maintain health and productivity is the cornerstone of the selective treatment paradigm (Bath and Wyk, 2001). Low-level parasitism is well tolerated by healthy hosts, and it stimulates natural immunity. Further, total elimination of parasites from grazing animals is an impractical and unsustainable goal.
Eggs per gram (EPG)
Individual FEC for Goat Herd 9000 8000 7000 6000 5000 4000 3000 2000 1000 0
67% of goats 20% of eggs
1 5 9 13 17 21 25 29 33 37 41 45 49 53 57 Individual FEC Animal Number
Figure I: Plot of individual fecal egg count results for a herd of 59 goats. The area to the right of the vertical line shows the FECs for the highest shedding goats – 1/3 of the herd, shedding 80% of the eggs.
with an effective anthelmintic. Animals scoring as a 1 or 2 are not anemic and therefore do not need treatment. Animals that receive a FAMACHA© score of 3 should be handled according to prevailing circumstances. For instance, young animals, and animals in herds or flocks where the majority of the animals have anemic scores and/or poor body conditions should be treated, particularly during periods of high Hc transmission (warm, moist conditions). Since many small ruminants that score as a 3/5 are not anemic, more animals will be treated than actually need it using the FAMACHA© system. However, the main benefit is that far less animals will be treated than when nonselective methods are used, so Refugia is maintained. FAMACHA© examinations should be conducted every 2-3 weeks, depending on the time of year and overall health of the herd. It must be emphasized that the FAMACHA© system is only applicable to management of Haemonchus contortus, and not all other internal parasites.
II. FAMACHA©: The FAMACHA© system takes advantage of GIN over-dispersion in a herd or flock, as well as anemia being the most prominent result of Hc infection. Anemia results in pallor of the conjunctiva and mucous membranes. This association was used by Dr Francois (“Fafa”) Malan to develop the FAMACHA© system for sheep and goat production systems in Africa (Malan, Wyk, et al, 2001). FAMACHA© is an acronym for FAfa MAlan’s CHArt. The FAMACHA© score is highly correlated with haematocrit (packed cell volume), and the fecal egg count in small ruminants and camelids with haemonchosis. The FAMACHA© system was validated for small ruminants in the United States in 2004 (Kaplan, Burke, et al, 2004) and camelids (Williamson, Storey, et al, 2009). The FAMACHA© evaluation is most accurate when performed in natural sunlight. The globe of the eye is gently retropulsed through the upper eyelid with one finger, and the lower eyelid everted so that the membrane color is visualized (See photo I). The color of the lower eyelid conjunctiva is compared with the FAMACHA© chart over a 1-3 second period, and assigned a score. The laminated FAMACHA© card has 5 scores based on the redness of the eyelid conjunctiva: 1 = deep red (non-anemic), 2= red-pink (non-anemic), 3 = pink (mild anemia), 4 white-pink = anemic, and 5 = white (severely anemic). Both eyelids are scored. If variation is noted between the 2 eyelid scores, the more anemic score is used in order to err on the side of safety. Animals with haemonchosis that score as a 4 or 5 should be treated
Photo I: Technique for exposing lower conjunctiva for FAMACHA© scoring (©2013 Bob Storey).
For the producer, the DrenchRite® Assay answers two critical questions related to a parasite management program in his herd/flock: 1) What GIN parasites infect my herd/flock? and 2) Which drugs are effective against the parasites on my farm? All that is required is proper sample collection and overnight shipping of the feces to the laboratory. Information, pricing, and instructions for submitting a sample for DrenchRite® analysis can be obtained by contacting Dr. Kaplan’s lab at (706) 5420742.
Photo II: Determining FAMACHA© eye score by comparing color of lower conjunctiva to FAMACHA© card (©2011 Bob Storey).
III. DrenchRite® Larval Development Assay (DrenchRite®). The DrenchRite® is an in vitro assay for the determination of anthelmintic efficacy (HorizonTechnology, 1996) on GIN of sheep, goats, and exotic hoof stock. It is performed in the laboratory of Dr. Ray Kaplan, Department of Infectious Diseases, University of Georgia College of Veterinary Medicine. The DrenchRite® utilizes eggs isolated from a composite fecal sample from a number of infected animals in a herd. Aliquots of the isolated eggs are placed into the wells of the DrenchRite® 96 well assay plate. The assay plate wells contain increasing concentrations of each class of anthelmintic; benzimidazoles (e.g., fenbendazole, albendazole), membrane depolarizers (e.g. levamisole, morantel tartrate), and the avermectins (e.g. ivermectin, abamectin). Moxidectin (which is from the milbemycen class) sensitivity is extrapolated from behavior of larvae in the higher concentrations of the ivermectin wells (Kaplan, Vidyashankar, et al, 2007). At the end of the incubation period (7 days at 25º C), the plate is examined using a microscope. The parasites present are identified by species, and the stage of development for each larva is determined and compared to the larvae that develop in the control wells. This information is then used to make a determination of susceptibility, suspected resistance, or resistance for each of the three classes of dewormers currently available.
IV. Conclusion: Gastrointestinal parasites in sheep and goats are a fact of life. Also, in this day and age, anthelmintic resistance is a fact of life. Neither parasites nor anthelmintic resistance is going to go away. Extending the life of current and possible new anthelmintics by decreasing selection pressure for AR is now the most important challenge facing sheep and goat producers. There is currently only one new anthelmintic in the development/approval process Zolvix® (monapantel). The market availability date for this new drug, in the US, is currently unknown. At present, we have the drugs that we have. Reducing AR selection pressure as well as dealing with existing AR parasites requires good animal husbandry as well as intelligent strategies that take advantage of parasite biology and host physiology. These strategies, such as selective treatment and proper dewormer use, are necessary for the small ruminant farmer to survive in the era of anthelmintic resistance. Failure to adapt these common sense principles of parasite management will also lead to the rapid development of resistance to any new drugs that come to market. The days of the magic bullet (inexpensive dewormers that work) are over; however, through careful herd management, selective use of dewormers, and exploitation of host physiology and parasite biology, successful parasite management in your herd or flock can be achieved. References: Barger, I. A. (1985). "The statistical distribution of trichostrongylid nematodes in grazing lambs." International Journal for Parasitology 15(6): 645-649. Bath, G. F. and J. A. v. Wyk (2001). Using the FAMACHA© system on commercial sheep farms in South Africa. Dobson, R. J., et al (2012). "Preserving new anthelmintics: A simple method for estimating faecal egg count reduction test (FECRT) confidence limits when efficacy and/or nematode aggregation is high." Veterinary Parasitology 186(1–2): 79-92.
HorizonTechnology (1996). DrenchRite® Larval Development Assay Standard Operating Procedures. Roseville, NSW, Australia, Horizon Technology Pty. Limited. Howell, S. B., et al (2008). "Prevalence of anthelmintic resistance on sheep and goat farms in the southeastern United States." Journal of American Veterinary Medical Association 233(12): 1913-1919. Kaplan, R. M., et al (2004). "Validation of the FAMACHA(C) eye color chart for detecting clinical anemia in sheep and goats on farms in the southern United States." Veterinary Parasitology 123(1/2): 105-120. Kaplan, R. M., et al (2007). "A novel approach for combining the use of in vitro and in vivo data to measure and detect emerging moxidectin resistance in gastrointestinal nematodes of goats." International Journal for Parasitology 37(7): 795-804.
Bob Storey is assistant research scientist and laboratory manager for the parasitology research labs of Dr. Ray M. Kaplan at the University of Georgia College of Veterinary Medicine. Bob also serves as the manager of the North American FAMACHA©program. Bob left his job with a Fortune 500 company to study veterinary technology, receiving both his VT degree as well as his RVT license (in Georgia) in 2007. Since then, he has authored or co-authored two dozen professional publications, made over 75 presentations at veterinary/professional conferences, trained hundreds of extension agents and producers in FAMACHA and SmartDrenching, and continued to maintain a small herd of Toggenburg goats and one llama. Bob’s research includes validating the use of FAMACHA© for South American camelids with Dr. Lisa Williamson; assessing the prevalence of Mycoplasma haemolamae in SAC in the Southeastern US with Dr. Alessandra Peligrini; and studying the pathology of Haemonchus contortus in Georgia’s SAC with Dr. Kaori Sakamoto. Bob’s recent activities include invited presentations at the 2015 Congress entitled “What Works with Worms,” in Pretoria, South Africa; at the 12th annual meeting of the American Consortium for Small Ruminant Parasite Control (ACSRPC); and at the 60th annual meeting of the American Association of Veterinary Parasitologists (AAVP). Bob has also given numerous FAMACHA workshops, including for the UGA Extension Service sponsored “Master Goat Producer” program.
Malan, F. S., et al (2001). "Clinical evaluation of anaemia in sheep: early trials." Onderstepoort Journal of Veterinary Research 68(3): 165-174. USDA-NAHMS (2011). Sheep 2011. Williamson, L. H., et al (2009). Evaluation of the FAMACHA© System in South American Camelids. World Association for the Advancement of Veterinary Parasitology, Calgary, Canada.
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Mary and John Bell Windy Hill Farm NC 844 Henderson Rd Tryon, NC 28782 828-894-3020 firstname.lastname@example.org www.happyalpacas.com
Sharon and Raymond Bogenschutz Georgian Oaks Farm 1973 Mary L. Trail Powder Oaks, GA 30127 678-427-2770 email@example.com www.openherd.com/farms/1005/ georgian-oaks-farm
Kathy Albert Heartland “Criations” Alpacas 2512 Knox Road 500 East Rio, IL 61472 309-368-7354 firstname.lastname@example.org www.hcalpacas.com Michelle Alexander Dakini Suri Alpacas 60375 Arnold Market Rd Bend, OR 97702 541-678-3306 email@example.com http://surimarket.surinetwork.org/ farms/4806 Michael and Sherry Alpert Awesome Acres Pacas & Pyrs 11800 S. Hiwassee Rd Oklahoma City, OK 73165 405-990-8205 firstname.lastname@example.org www.pacasnpyrs.com Carl & Regina Alvaraz Braecroft Suri Alpacas PO Box 26 Mayhill, NM 88339 575-687-3697 email@example.com surimarket.surinetwork.org/ farms/2928 Patti Anderson and Alan Anderson Wild Rose Suri Ranch 3623 Harmony Church Rd Havre de Grace, MD 21078-1015 410-734-7084 firstname.lastname@example.org www.WildRoseAlpacas.com
Roy and Rex Anderson Anderson Acres Alpacas, LLC 8812 S 230 Plaza Circle Gretna, NE 68028 402-740-2711 email@example.com www.openherd.com/farms/4899/ anderson-acres-alpacas-llc
Linda Bader and Cindy Smith Shady Hollow Suri Alpacas 4810 McMillan Road Bad Axe, MI 48413 989-658-8629 firstname.lastname@example.org www.shsurialpacas.com
James Bell and Carmela Osborne Bellborne Farm 835 South Main Street Findlay, OH 45840 419-348-0973 email@example.com www.bellbornefarm.com Kim Bisceglia McCarty Creek Ranch P.O. Box 2216 Flournoy, CA 96029 530-833-5431 firstname.lastname@example.org www.openherd.com/farms/3860/ mccarty-creek-ranch
Barbara Boineau River Run Ranch 1706 S Riverbend Rd Wanship, UT 84017 435-901-1274 email@example.com www.RiverRunAlpacas.com Randy and Beth Brealey Chelsea Farms 19450 208th Ave SE Renton, WA 98058 206-229-8845 firstname.lastname@example.org www.TheAlpacaPlace.com
Brian and Katie Barfield Silverfield Farm 34203 E Lacomb Road Lebanon, OR 97355 720-530-5648 email@example.com
Karen Biscella Victoria Lane Alpacas 48216 Metz Rd New Waterford, OH 44445 330-503-9561 firstname.lastname@example.org www.victorialanealpacas.com
Beverly and Jason Brehm Rogue Suri Alpacas 1075 Washburn Lane Medford, OR 97501 541-292-6844 email@example.com
William E. Barnett, DVM Alpacas of America, LLC 16641 Old Highway 99 SE Tenino, WA 98589 360-264-7585 firstname.lastname@example.org www.alpaca1.com
Dean and Connie Blechle Breezy Ridge Alpacas LLC 908 PCR 810 Perryville, MO 63775 573-547-2217 email@example.com www.breezyridgealpacas.com
Bill and Carol Barrett Heartsease Alpacas LLC 7851 N. Red Hill Rd Ellettsville, IN 47429 812-876-5228 firstname.lastname@example.org www.heartseasealpacas.com
Bob and Vicki Blodgett Suri Land Alpaca Ranch 10371 N 2210 Rd Clinton, OK 73601 641-831-3576 email@example.com www.alpacanation.com/suriland.asp
Sandra and Clint Brown Enchanted Grove Alpacas Box 1138 Portage, Manitoba Canada R1N 3J9 204-297-7329 firstname.lastname@example.org www.enchantedgrovealpacas.ca
The Suri Network membership year now runs from June 1 to May 31 of the following year for all members. All members current in their dues as of June 1, 2016 are included in this listing. Members are listed alphabetically by last name. All members have the ability to update their contact information on-line at any time through their account on the Suri Network website. We encourage members to keep their contact information complete and up to date, so that potential clients can easily reach them. Should you need assistance updating your information, feel free to contact the office at (970) 586-5876 and they will be happy to assist you. The Suri Network is not responsible for, and expressly disclaims all liability for, damages of any kind arising out of use, reference to, or reliance on any information contained within this directory.
Suri Network Membership Directory Dawn Browning and Susan Feil Sie Sutter Suri Alpaca, LLC 703 Fort Hill Rd Maysville, KY 41056 859-338-2946 email@example.com www.openherd.com/farms/3829/siesutter-suri-alpaca-llc Bill and Paula Buchner Alpacas of Greater Tennessee 10925 Hwy 58 Georgetown, TN 37336 423-344-5252 firstname.lastname@example.org www.alpacasofgreatertn.com Judith Buning Silver Lining Suris 1150 W. Brown Rd Mayville, MI 48744 810-656-7784 email@example.com www.slsuris.com Willard Burney Sound of Freedom Farm, LLC 3936 Charity Neck Road Virginia Beach, VA 23457 844-763-3276 firstname.lastname@example.org www.soundoffreedomfarm.com Kellie and David Butsack Noble Glen 555 Valley Road Mohrsville, PA 19541 610-926-9690 email@example.com www.nobleglen.com Jay Call and Gina White Finca Solaz 8100 FM 2325 Wimberley, TX 78676 512-847-2381 firstname.lastname@example.org Kathleen and Tom Callan Zena Suri Alpacas 35401 S 580 Rd Jay, OK 74346 804-389-2579 email@example.com www.zenasurialpacas.com Dawn Callaway and Brian Stole BDS Ranch 6301 East Mountain View Road Paradise Valley, AZ 85253 480-529-1984 firstname.lastname@example.org
Albert J. and Rebecca A. Camma The Alpacas of Spring Acres 3370 Big B Rd Zanesville, OH 43701 740-796-2195 email@example.com www.thealpacasofspringacres.com Gail Campell Ameripaca Alpaca Breeding Co. P.O. Box 256 Galesville, MD 20778 410-867-4204 firstname.lastname@example.org www.ameripaca.com Jeanette and James Campbell Alpacas at Willowbrook Farm 24365 Pflumm Road Louisburg, KS 66053 913-879-2066 email@example.com www.alpacasatwillowbrook.com Debbie and Ed Chesna Ledge View Alpaca Farm W 4173 County Road H Fond Du Lac, WI 54937-9653 920-923-6925 firstname.lastname@example.org
Deb and Doug Christner Akuna Matada Suri Alpacas 28444 Redlands Mesa Rd Hotchkiss, CO 81419 970-872-4446 email@example.com
Mary Lou Clingan Waywood Farm Alpacas 14136 Township Road 108 Findlay, OH 45840 419-306-2512 firstname.lastname@example.org Barbara and Randy Coleman Wings & A Prayer Alpacas 18100 S. Hwy. 99W Amity, OR 97101 503-310-9367 email@example.com www.wingsandaprayeralpacas.com Lisa Collura and Robert Figular Memory M-Acres 33 New Rd Lambertville, NJ 08530 609-902-3551 firstname.lastname@example.org www.memorymacres.com
Gregory Conley 3410 N Placita Herradura Douglas, AZ 85607 609-680-9203 email@example.com Michelle Cooper Cooper Now Alpacas 4213 Arkney Court Fort Wayne, IN 46814 260-615-1721 firstname.lastname@example.org Dan and Cari Corley Alta Vida Alpacas 9535 E. Parker Rd Parker, CO 80138 303-884-7374 email@example.com www.altavidaalpacas.com John and Nikki Creasey White Rock Alpacas 5240 Garton Rd Castle Rock, CO 80104 713-494-2867 firstname.lastname@example.org Heather Cross Suri Charisma Alpacas 18575 Ramah Hwy Ramah, CO 80832 719-541-0321 email@example.com R.T. Crowe, II Bar C Ranch 211 Slim Buttes Rd Chadron, NE 69337 775-750-0099 firstname.lastname@example.org
Brenda and Frank Crum Golden Spirit Alpaca Ranch 17902 Spencer Rd Odessa, FL 33556 727-809-2253 email@example.com www.goldenspiritalpaca.com Lynda Cunningham and Jeff Bennett Serenity Valley Alpacas 2026 Waring St Seaside, CA 93955-3215 831-869-0751 firstname.lastname@example.org www.sv-alpacas.com
Dale and Jan Davis Derwydd Alpacas 24485 Derwydd Way Esparto, CA 95627 530-908-3534 email@example.com www.derwyddalpacas.com Dave DeGroot DDF Alpacas PO Box 510 Odell, OR 97044 541-806-6262 firstname.lastname@example.org Hillary Devin and Scott Phillips Shambalah Alpaca Ranch 7157 E Park Dr Franktown, CO 80116 303-588-2076 email@example.com www.shambalahalpaca.com Diane and Bobby Dickerson Rock’n D Enterprises 11550 Hunt Lane Guthrie, OK 73044 405-850-6533 firstname.lastname@example.org www.rockndenterprises.com Kim Dietz KaJota Meadows LLC 1835 Risser Mill Rd Mount Joy , PA 17552 717-492-4114 email@example.com www.kajotameadows.com Tracy DiPippo Angel Dreams Alpacas 291 Race Street Grass Valley, CA 95945 805-432-9344 firstname.lastname@example.org www.Alpacasdream.com Evi Dixon Spanish Peaks Alpacas LLC 3180 Curtis Lane Manhattan, MT 59741 406-579-9694 email@example.com www.sp-surialpacas.net Jack and Miriam Donaldson Alpaca Jack’s Suri Farm 13939 CR 215 Findlay, OH 45840 419-423-3890 firstname.lastname@example.org www.alpacajack.com
Suri Network Membership Directory Tilly and Katie Dorsey DAFI Alpacas P.O. Box 55 Butler, MD 21023 410-591-0691 email@example.com www.dafi.com
Lucy Lee and King Fowler LunaSea Alpaca Farm 11812 Ridge View Circle Clermont, FL 34711 352-223-9457 firstname.lastname@example.org www.lunaseaalpacafarm.com
Kathy Graziani Flame Pool Alpacas, Ltd. 9124 Flamepool Way Columbia, MD 21045 410-884-5397 email@example.com www.flamepoolalpacas.com
Lisa Haselhorst Grandview Suri Alpacas 12102 N First St Parker, CO 80134 303-910-4975 firstname.lastname@example.org www.grandviewsurialpacas.com
Victor Leland Epperson Moon Shine Alpacas 297 Bushnell Rd Douglas, GA 31533 530-518-4094 email@example.com www.moonshinealpacas.com
Lona Nelsen Frank and John Frank ALPACAS of Tualatin Valley, LLC 22750 SW Rosedale Rd Beaverton, OR 97007 503-649-2128 firstname.lastname@example.org www.alpacatv.com
Collins and Nikki Griffith Sandollar Alpacas 2001 S Washington St Kennewick, WA 99337 509-378-5833 email@example.com www.sandollaralpacas.com
Britt and Patty Hasselbring Hasselbringâ€™s Harmony Ranch PO Box 630 Concordia, MO 64020 816-769-3939
Norm Evans, DVM Florissant, MO
Chris and Jess Fredericks Whistling Pines Ranch 499 4 1/2 Ave Clayton, WI 54004 715-419-0127 firstname.lastname@example.org www.whistlingpinesranch.com
Jennifer Hack Triple H Ranch 4098 N Hwy 67 Sedalia, CO 80135 720-733-2672 email@example.com www.triplehalpacas.net
Brenda Hauck Top of the Ridge Ranch, LLC 9990 Hahn Rd Calhan, CO 80808 720-244-6075 firstname.lastname@example.org www.topoftheridgeranch.com
Carol Furman Carrageen Alpacas 82 West Gray Road Gray, ME 04039 207-653-1144 email@example.com www.carrageensurialpacas.com
Steve and Laura Hall BuzznHum Alpacas LLC 15851 NW Willis Road McMinnville, OR 97128 503-434-6358 firstname.lastname@example.org www.buzznhumalpacas.com
Mike and Mary Hayden Quest Alpacas 20911 Ehle Rd Woodburn, IN 46797 260-632-4399 email@example.com www.QuestAlpacas.com
Jerad Gan The Alpaca GANg 1174 Conoco Rd Mulberry Grove, IL 62262 618-960-6437 firstname.lastname@example.org www.thealpacagang.com
Rebecca Hammer Orchard Hill Alpacas PO Box 842 Elkins, WV 26241 304-636-5519 email@example.com
Joy Hays SuperFleece Alpacas, LLC 150 S Harrison St, Unit 5 Denver, CO 80209 913-481-5243 firstname.lastname@example.org
Cheryl Gehly Cria-tivity Alpacas 161 West Shore Road B-11 Warwick, RI 02889 508-404-4373 email@example.com
Ron and Kathy Harelstad R K Ranch 626 18th Street Chetek, WI 54728 715-837-1759 firstname.lastname@example.org
Louise and Robert Hazen Blue Mesa Alpacas, LLC 10 Blue Mesa Rd Santa Fe, NM 87508 505-473-9316 email@example.com www.BlueMesaAlpacas.com
Scott and Laurie Findlay Alpacas of El Dorado 4535 Boo Bear Lane Somerset, CA 95684 530-642-8082 firstname.lastname@example.org www.alpacasofeldorado.com Ben and Lynda Fisco Humming Hill Suri Farm Ltd. 12100 Pekin Road Newbury, OH 44065-9622 440-564-5114 email@example.com www.humminghill.com Carrie and Scott Fletcher Cabin Critters 9779 Beaver Pond Rd Conifer, CO 80433 303-718-1597 firstname.lastname@example.org
June Ford Shady J Ranch PO Box 2737 Ramona, CA 92065 858-531-6696 email@example.com Robert and Debra Fordam Miners Mountain Alpacas PO Box 135 Teasdale, UT 84773 435-680-7687 firstname.lastname@example.org
Stacey Goss Goldyn Rae Alpacas PO Box 2004 Elizabeth, CO 80107 303-819-8001 email@example.com www.goldyn-rae-alpacas.com Bob Graf Alpaca Bob 611 S Palm Canyon Dr #7452 Palm Springs, CA 92264 760-327-7980 www.AlpacaBob.com
Cindy Harris Alpacas at Windy Hill 7660 Bradley Road Somis, CA 93066 805-907-5162 firstname.lastname@example.org www.alpacalink.com
Karl and Janice Heinrich Long Hollow Suri Alpacas 698 Wallace Road Gallatin, TN 37066 615-452-7852 email@example.com www.longhollowalpacas.com Nancy and David Helwig 925 Sterling Alpacas 10451 Valley Drive Plymouth, CA 95669-9515 209-245-3438 firstname.lastname@example.org www.925suris.com
Suri Network Membership Directory Heather Hernandez Joya de Suri 6920 6150 Rd Olathe, CO 81425 970-323-6835 email@example.com Leslie Herzog Herzog Alpacas, LLC 1762 McCraren Rd Highland Park, IL 60035 847-702-7731 firstname.lastname@example.org
Roxann and Jay Hirst Crystal Butte Farm 8330 316th Pl SE Issaquah, WA 98027 703-687-7676 email@example.com Jim and Jane Olson-Holzer
Unisource Suri Alpacas & Llamas, LLC
144 Osprey Circle Hope, ID 83836-9704 208-264-5101
Susan E. Inman Flights of Fancy Farm 61750 Cougar Trail Bend, OR 97701-9636 541-598-7417 firstname.lastname@example.org Amber Isaac Silken Suri Alpaca Ranch 6219 Lake Gulch Rd Castle Rock, CO 80104 303-258-7835 email@example.com www.silkensuri.com Denise and Jeff Johnson Zephyr Hill Farm 10297 7 Mile Rd Evart, MI 49631 248-770-5514 firstname.lastname@example.org www.openherd.com/farms/7100/ zephyr-hill-farm Julie Johnson and Tim Gallagher Cannon River Alpacas 31536 Cannon River Blvd Northfield, MN 55057 507-366-1208 email@example.com
Dianna and Jack Jordan Alpacas of Somerset Farm PO Box 190 Somerset, CA 95684 530-620-6033 firstname.lastname@example.org www.alpacasofsomersetfarm.com Nicholas Judson Slick and Twisted Suri Alpacas Ltd 7083 S Knolls Way Centennial, CO 80122 303-258-7835 email@example.com Ann and Tony Kaminski Break Loose Farm 5233 Hanover Pike Manchester, MD 21102 410-374-4383 firstname.lastname@example.org www.breakloosefarmalpacas.com Pamela Kelly Bridgetown Suri Alpacas 9875 Bayside Rd Machipongo, VA 23405 757-709-0400 email@example.com www.bridgetownsurialpacas.com Susan King Big Timber Alpacas 29400 SW Heater Rd Sherwood, OR 97140 503-799-6941 firstname.lastname@example.org www.bigtimberalpacas.com Vern and Mary Knadler Big K’s Suri Alpacas 605 Rex Church Road Saint Paul’s, NC 28384 910-635-9010 email@example.com www.bigkalpacafarm.com Steven, Rose Ann and Tasha Knoblock
Knoblock’s Prairie Ranch 179 270th Street Sabetha, KS 66534 785-284-2589 firstname.lastname@example.org www.knoblocksalpacas.com
Linda K. Kondris Pines Edge Suri Alpacas 12245 Melba Rd Black Forest, CO 80106-8966 719-495-9633 email@example.com www.pinesedge.com
Judith A. Korff LadySong Farm 2473 Bunker Hill Rd Randolph, NY 14772 716-499-0383 firstname.lastname@example.org
Margit Korsak Boulder Hill Alpacas 315 Merkle Rd Boyertown, PA 19512 610-367-8486 email@example.com www.boulderhillalpacas.com Karen Kovisto Symphony Fibers 6670 Sunset Circle Kiowa, CO 80117 818-326-7393 firstname.lastname@example.org www.SymphonyFibers.com Celeste Kreiensieck and Bob Cross Thunder Mountain Alpacas 21518 NE 192nd Ave Battle Ground, WA 98604 360-666-4553 email@example.com www.ThunderMountainAlpacas.com Lorrie Lake Loving Life Alpacas 34706 Minesinger Trail Polson, MT 59860 406-261-8772 firstname.lastname@example.org www.openherd.com/farms/4444/ loving-life-alpacas Darlene Lander Dun Roving Farm 359 Dodge Rd Frewsburg, NY 14738-9742 716-569-2198 email@example.com www.dunrovingfarm.com Mary-Margaret Lannon Percussion Rock Alpaca Farm 6669 N Sioux Chicago, IL 60646 773-577-0217 firstname.lastname@example.org
Kirk and Julie Lentz Alpine Vista Suri Alpacas LLC 73 Wineglass Loop South Livingston, MT 59047 406-222-0472 email@example.com www.alpinevistasurialpacas.com Nancy Lindemood 2 Point Farm, LLC 6330 Warsaw Rd Dry Ridge, KY 41035 859-428-9220 firstname.lastname@example.org www.2pointfarmalpacas.com Margaret Long and Susan Simonton
Little Gidding Farm Suri Alpacas 17585 Eagle Ave Lester Prairie, MN 55354-7831 320-224-4204 l email@example.com www.lgfsuris.com Suzi Mancuso Ponderosa Paca Farm 20 Natures Crossing Middleboro, MA 02346 508-989-7961 firstname.lastname@example.org Cora and Randy Marburger Next Chapter Alpacas 801 Middle Cove Drive Plano, TX 75023 214-395-8845 email@example.com www.nextchapteralpacas.com
Anita and Richard Marlin Acappella Junction Alpacas 502 Ostwalt Amity Road Troutman, NC 28166 919-599-7359 firstname.lastname@example.org www.accappellajunctionalpacas.com Jackie Mathieson Alpaca Road, LLC 1135 Camp Road Denton, MD 21629 410-241-4367 email@example.com www.alpacaroad.com Teresa and Joe Maxwell Stone Spring Suri Farm 18 McHenry Rd Cochranville, PA 19330 610-593-6694 firstname.lastname@example.org www.stonespringsuri.com
Suri Network Membership Directory Curtis May Sweet Escape Alpaca Ranch 6442 Quartz Cir Arvada, CO 80007 303-908-7112 email@example.com
Bryan and Becky Miltenberger Hidden Creek Alpacas 33347 East Punkin Center Rd Hermiston, OR 97838 541-571-0394 firstname.lastname@example.org www.openherd.com/farms/4788/ hidden-creek-alpacas
Ann Mayes Alpacas d’Auxvasse 9266 County Rd 1012 Auxvasse, MO 65231 573-386-3462 email@example.com www.alpacasauxvasse.com
David Moran and Lori Wall Crimson Shamrock Ranch PO Box 7 Eglon, WV 26716 304-735-6413 firstname.lastname@example.org www.crimsonshamrockalpacas.com
Alvina Maynard River Hill Ranch 680 River Hill Rd Richmond, KY 40475 859-408-5132 email@example.com www.riverhillranch.us/
Jim and Jean Morgan J4 Alpacas 7711 N Valley Hill Road Woodstock, IL 60098 815-759-0247 firstname.lastname@example.org www.j4alpacas.com
Jill M. McElderry-Maxwell Bag End Suri Alpacas of ME, LLC 226 Snakeroot Rd Pittsfield, ME 04967 207-660-5276 email@example.com www.bagendsuris.com
Sandra and Ralph Muraca Misty Mountains Accoyo Suris 121 Stoners Hill Road Raphine, VA 24472 540-377-2110 firstname.lastname@example.org www.mmasalpacas.com
Candy and Ray McMahan and Joe Miller Miller - McMahan Alpacas, LLC 1897 Ashland Rd Ruffin, NC 27326 402-580-5453 email@example.com www.mmalpacas.com
Kent and Sandy Murray Lizard Hill Suri Alpacas 351 Purdy Mesa Road Whitewater, CO 81527 970-243-3520 firstname.lastname@example.org
Becky and Tom McMillan Magic Willows Alpacas 6340 Arthur Rd Hartford, WI 53027 414-217-5836 email@example.com www.magicwillowsalpacas.com Thomas and Collene Miller Purdy Lil’ Suri Alpaca Farm, LLC 9454 Taylorsville Rd Huber Heights, OH 45424 937-233-3509 firstname.lastname@example.org www.purdylilsuri.com/
Richard and Sharon Musser Shardick Suri Farm 54 Old State Rd Jonestown, PA 17038 717-865-1809 email@example.com https://www.openherd.com/ farms/4275/shardick-suri-farm Jennifer Myka Free Radical Ranch 1103 Bracht-Piner Road Morning View, KY 41063 859-462-2344 firstname.lastname@example.org www.freeradicalranch.com
Richard and Leanne Nakashima Eldora Suri Alpacas 31941 Hamilton Creek School Road Lebanon, OR 97355-9729 720-840-6585 email@example.com www.eldorasurialpacas.com
Narvel and Debbie Pettis Sweet Blossom Alpaca Farm 37543 Pappy Road Dade City, FL 33523 813-335-7387 firstname.lastname@example.org www.sweetblossomalpacas.com
Diane and Julian Nicholson Nicholson Alpacas and Llamas P.O. Box 5378 Twin Falls, ID 83303 208-734-5917
Charlene and Russ Piar Thunder Mile Ranch 10879 State Hwy M Wright City, MO 63390 636-544-2200 email@example.com www.thundermileranch.com
Onneke O’Brien SuCaya Farms 576 Lake View Dr Smithfield, ME 04978 207-634-2189 firstname.lastname@example.org
Judy Olukotun Pear Tree Farm 125 Hopewell-Wertsville Rd Hopewell, NJ 08525 609-466-9621 email@example.com www.openherd.com/farms/2905/ pear-tree-farm
Karina and Michael Pomroy Peruvian Link Co. 589 Airline Rd Amherst, ME 04605 207-584-3200 firstname.lastname@example.org www.peruvianlink.com Michelle Pressler and William Ward
iMpress Alpacas 14036 Clover Rd Rockton, IL 61072 815-713-5234 email@example.com www.impressalpacas.com
Dennis and Christy Pace Pacesetter Alpacas 5546 W Plymouth Church Road Beloit, WI 53511 608-879-2770 firstname.lastname@example.org www.pacesetteralpacas.com
Teri and Kraig Quamme Red Gate Alpaca Farm 11751 Dundas Blvd Dundas, MN 55019 612-501-9161 email@example.com www. redgatealpacafarm.com
Tracy Pellegrino Faith & Fleece Alpacas 2664 N Mountain Ave Upland, CA 91784 626-716-0943 firstname.lastname@example.org www.faithandfleece.com
Brenda Quinones Natural People Ranch Suri Alpacas PO Box 92 Palmer Lake, CO 80133 719-287-6726 email@example.com
Marlene and Jonathan Peltier Mystical Acres Alpacas 9678 N Co Road, 25-A Sidney, OH 45365 937-492-0776 firstname.lastname@example.org www.mysticalalpacas.com
Ken and Claudia Raessler SuriPaco LLC PO Box 1477 Yarmouth, ME 04096-2477 207-712-5833 email@example.com
John and Jan Rager Silver Threads Alpaca Ranch 298 L’Esprit Farm Rd LaGrange, KY 40031 502-222-2238 firstname.lastname@example.org www.silverthreadsalpaca.com
Suri Network Membership Directory Sheri Raterink Dream Seeker Suris 9856 Federal Rd Howard City, MI 49329 email@example.com Allen and Becky Rebman and Richard, Carol and Andrew Reed Over Home Alpacas LLC 18 Midway Rd Bethel, PA 19507 610-488-1355 firstname.lastname@example.org www.overhomealpacas.com Leslie and Scott Rebtoy Healing Springs Suris PO Box 1400 Westville, OK 74965 918-629-2840 HSSsuris@gmail.com www.Healingspringssuris.com Jane and Ron Reed Suri Haven 7940 W Barrington Rd Kirkland, AZ 86332 402-314-1323 email@example.com www.surihaven.com Kahley Reps Reps Family Farm 113 Renea Dr St Charles, MN 55972 507-254-2327 firstname.lastname@example.org Doug and Julie Rice R&R Suris 11110 Kubon Rd Montague, MI 49437 231-893-7208 email@example.com www.AlpacaNation.com/ R&Ralpacas.asp Marc and Nanci Richards March Hare Station 1329 County Road 2400 E Saint Joseph, IL 61873 217-469-2767 firstname.lastname@example.org
Julie and Ted Ritschard B I Bar Ranch 8720 Moss Rock Rd Colorado Springs, CO 80908 719-495-1279 email@example.com bibarranch.com
Kate and Ed Robie Ash Hill Alpacas 1605 Carpenter Pike Versailles, KY 40383 404-441-3155 firstname.lastname@example.org https://www.facebook.com/ ashhillalpacas/ Janet and Bob Rodgers Rodgersâ€™ Reserve Alpaca Farm 400 E. Adario W. Rd Greenwich, OH 44837 419-895-9922 email@example.com www.openherd.com/rodgersreserve Christine Rogers Whisper Meadows Alpacas 4451 Whisper Lane DePere, WI 54115 920-337-0646 firstname.lastname@example.org www.alpacanation.com/ whispermeadows.asp Amy and Dave Self Canyon Country Suri Alpaca Ranch 12500 Bostwick Park Rd Montrose, CO 81401 970-765-6744 email@example.com surimarket.surinetwork.org/farms/4970
Tim and Beth Sheets Heritage Farm Suri Alpacas 4175 N 1200 W Flora, IN 46929 765-566-3077 firstname.lastname@example.org www.ourheritagefarm.com Sheryl and Sheldon Shenk Hay Creek Station PO Box 589 Florissant, CO 80816 719-689-6666 email@example.com www.yoursuriconnection.com Laurel R. Shouvlin Bluebird Hills Farm 3617 Derr Road Springfield, OH 45503 937-206-3936 firstname.lastname@example.org www.bluebirdhills.farm
Stacy Siler USS Alpacas PO Box 671 Wheaton, MO 64874 417-342-6170 email@example.com June Sims Cloud Dancing Alpacas 1365 Long Rd Defuniak Springs, FL 32433 850-758-1270 firstname.lastname@example.org Gary and Michele Siplivy Suri Alpacas of Shiloh Farm 278 Old State Rd Shermans Dale, PA 17090 717-514-1819 email@example.com www.shilohsurialpacas.com Cindy and Jay Smith Weeping Willow Farm LLC 1265 Cherokee Dr De Leon Springs, FL 32130 386-277-2043 firstname.lastname@example.org
Michael and Anita Smith The Triple Z Alpaca Farm 9100 N 000 Rd Decatur, IN 46733 260-724-4809 email@example.com www.triplezalpacas.com Valarie Smith Double Diamond Ranch 215 Quail Ridge Road Amarillo, TX 79118 806-729-6113 firstname.lastname@example.org Kristie and Brion Smoker Sweet Valley Suris 5701 Valley Glen Road Annville, PA 17003 717-867-2897 email@example.com www.sweetvalleysuris.com Brad and Jandy Sprouse Great Lakes Ranch 5718 S Bohemian Rd Maple City, MI 49664 231-228-3859 firstname.lastname@example.org www.greatlakesranch.com
Loren and Judy Stevens Stevens Llama Tique & Suri Alpacas 1449 Red Canyon Rd Canon City, CO 81212 800-469-5262 email@example.com www.stevenstique.com Rick and Kathy Stumpf Prairie Lake Alpacas 13711 John Kline Rd Smithsburg, MD 21783 301-416-0833 firstname.lastname@example.org www.prairielakealpacas.com Kim Taha Taha Suri Alpaca Ranch, LLC 23109 US Highway 40 Golden, CO 80401 303-704-7928 email@example.com www.tahaalpacas.com Keiko Takimoto-Makarczyk and Matt Makarczyk Wisteria Suri Ranch 875 County Road 454 Taylor, TX 76574 512-856-2467 firstname.lastname@example.org www.wisteriasuriranch.com Victoria Telesko Love Me Alpacas 23253 Cty Rd X Kiel, WI 53042 920-797-9096 email@example.com www.LoveMeAlpacas.com Susan Tellez
Camelid Alliance - Resource Consulting
3195 Dowlen Rd. #101-313 Beaumont, TX 77706 409-656-2140 firstname.lastname@example.org
David, Nancy and Nick TenHulzen Park View All American Alpacas 3001 SW Schaeffer Rd West Linn, OR 97068-9611 503-638-3692 email@example.com www.ParkViewAlpacas.com Ingrid Thomas Hadleigh Grange 3269 N Reed Station Rd Desoto, IL 62924-3417 firstname.lastname@example.org
Suri Network Membership Directory Kerry and Jodi Thompson Patriot Lane Alpacas 80189 Patriot Lane Hermiston, OR 97838 541-720-2194 email@example.com www.patriotlane.com Dana Tiedeman Riverside Suri Alpacas 33073 Lakeview Dr Lake City, MN 55041 651-380-6336 firstname.lastname@example.org www.riversidesurialpacas.com Andy and Cheryl Tillman Tillman Llamas & Suri Alpacas 20510 Swalley Rd Bend, OR 97701 541-389-1065 email@example.com www.tillmansranch.com Christie Tolj 24405 SW Ballston Rd Sheridan, OR 97378 503-006-4655 firstname.lastname@example.org Patty Toney Dixieland Alpaca Farm 376 Blue Door Rd Portland, TN 37148 615-325-6439 email@example.com www.dixielandalpacafarm.com Marcia Traudt M & J Alpaca Farm 32382 Hwy 14 Aurora, NE 68818 402-737-3307 firstname.lastname@example.org Gary and Cindy Truitt Weatherd’T Ranch, LLC 11021 County Road 102 Elbert, CO 80106 303-648-9228 email@example.com www.wtralpacas.com Kathy and Garry Umscheid Evergreen Acres Alpacas P.O. Box 117 Arden, MB Canada R0J 0B0 204-368-2467 firstname.lastname@example.org www.evergreenacres.org
Liz and Chris Vahlkamp Salt River Alpacas/North American Suri Co. 7200 Waterman St. Louis, MO 63130 314-440-1627 email@example.com www.saltriveralpacas.com
Jim Weir Wildlife Ranch Suri Alpacas 10500 Wildlife Way Littleton, CO 80125 303-885-3377 wildliferanchsurialpacas@gmail. com
Bill and Heather Vonderhaar Alpaca Bella Suri Farm - ABF 5455 Belwood Lane Morrow, OH 45152 513-899-2304 firstname.lastname@example.org www.alpacabella.com
Tim and Dana Welch Purgatory Falls Alpaca Farm 195 Purgatory Falls Road Lyndeborough, NH 03082 603-654-7690 email@example.com www.purgatoryfallsalpaca.com
Bob Wargowsky Windrider Suri Ranch PO Box 1331 Norwood, CO 81423 970-327-0149 firstname.lastname@example.org
Deb Wellinghoff Northern Prairie Alpacas, LLC 7470 Jakes Prairie Rd Sullivan, MO 63080 618-558-8390 email@example.com www.NorthernPrairieAlpacas.com
Gail and Paul Wasserstein Andean Vista Ranch 4101 Oak Ridge Road Crystal Lake, IL 60012 847-366-9147 firstname.lastname@example.org www.andeanvistaranch.com
Marilyn Wentworth Alpacas at Phoenix Hill Farm, LLC 8266 Rock Riffle Road Athens, OH 45701 740-591-7669 email@example.com www.alpacasatphonexhill.com Joyce and Greg White Tinkers Creek Alpacas LLC 3851 Sanford Rd Rootstown, OH 44272 330-524-2077 firstname.lastname@example.org
Diane Watson Sweet Heart Suri Alpacas 4813 Church St Conneaut, OH 44030 440-224-1868 email@example.com www.sweetsurialpacas.com Dennis and Rose Watts Thunder River Suri Alpacas 9627 Carlisle Road Dillsburg, PA 17019 717-994-4055 firstname.lastname@example.org www.thunderriveralpacas.com Brett and Donna Weeks Grey Meadows Alpaca Farm 1835 Underwood Road Gambrills, MD 21054 301-980-7019 email@example.com
Mary Wilcox High Country Alpacas LLC 3805 E Equestrian Trail Phoenix, AZ 85044-3008 480-296-8588 firstname.lastname@example.org www.hcaalpacas.com Mike and Janet Wilkins Wilkins Livestock LLC P.O. Box 7221 Star Valley Ranch, WY 83127 402-362-9223 email@example.com www.wilkinslivestock.com
Kathy and Joe Williams KJ’s Alpaca Ranch 7476 Shepler Church Ave SW Navarre, OH 44662 330-879-2483 firstname.lastname@example.org www.kjsalpacas.com Doug and Deanna Wilner Daydreamer Ranch Alpacas 16019 Green Road Harvard, IL 60033 815-943-7004 email@example.com
Dr. Gary Wilson The Midnight Moon Alpaca Ranch 937 Hillside Ave Elmhurst, IL 60126 630-921-0414 garyw@ themidnightmoonalpacaranch.com www.openherd.com/farms/2413/ the-midnight-moon-alpaca-ranch Livia Wright 6111 Irby Ln W Lakeland, FL 33811 863-937-7980 firstname.lastname@example.org Jessica Wyatt 2135 Burning Ridge Dr Franktown, CO 80116 505-263-6020 email@example.com Mary Yaros First Light Alpacas 3855 Paseo Del Prado Boulder, CO 80301 720-381-0132 firstname.lastname@example.org Melisa and Terry Yopp Berry Sweet Alpacas 12167 Centerpoint Church Road Prairie Grove, AR 72753 479-871-4304 email@example.com www.berrysweetalpacas.com Cheryl and Rick Yopp City Girl Alpaca 9 Greak Oak Court North Little Rock, AR 72116 501-753-8480 firstname.lastname@example.org
Suri Network Membership Rory York Mystic Winds Farm PO Box 36 Florence, CO 81226 719-429-4949 email@example.com www.MysticWindsFarm.com Barbara Zachary New Age Alpacas 9460 Santa Clara Road Atascadero, CA 93422 805-286-2070 firstname.lastname@example.org www.newagealpacas.com
Elizabeth Saletta Kiel, WI Marina Welch Tampa, FL Sarah Withrow Lees Summit, MO
Norm and Mary Zahn Coldwater Creek Alpacas 5254 Younger Rd Celina, OH 45822 419-678-8621 email@example.com www.ColdwaterCreekAlpacas.com Dave and Louann Zapicchi Hobble Hill Farm LLC 1322 Sylvan Rd Perkasie, PA 18944 215-795-0670 firstname.lastname@example.org www.hobblehill.com Barb and Glen Zimmerly Glenbar Alpacas 20801 NE 50th Ave Ridgefield, WA 98642 360-574-5428 email@example.com www.glenbaralpacas.com SURI ENTHUSIAST MEMBERS Sonja Boeff Arvada, CO Angela Echeverria Kansas City, MO Kathryn Felty Reston, VA Stacy Heydt Marshfield, MO Florence Morehead Wildwood Alpaca Farm Wellborn, FL Donna Rudd Clive, Alberta, Canada
2 Point Farm 23 Alpaca Embryo Technology, LLC 12 Alpacas of Oklahoma (Blastoff) 13 Ameripaca Alpaca Breeding Company 3 Bag End Suri Alpacas of Maine, LLC 13 Big Timber Alpacas 2 Experienced Suri Partners 44-45 Hasselbringâ€™s Harmony Ranch 67 Healing Springs Suris 65 Heritage Farm Suri Alpacas 33 J4 Alpaca Farm 52 KJs Alpaca Ranch 7 Long Hollow Suris/New Era Fiber 57 M&J Alpaca Farm 65 Nebraska Alpaca Company 8 Over Home Alpacas 23 Pines Edge Suri Alpacas 3 Pucara International 12 Raynay Alpaca Farm 8 Rogue Suri Alpacas
Salt River Alpacas 31 Sie Sutter Suris 13 Sweet Blossom Alpacas 65 Sweet Valley Suris 13
Photograph courtesy of Linda Kondris @2016