Blood Collection from Mice and Hematological Analyses on Mouse Blood.

Blood Collection from Mice and Hematological Analyses on Mouse Blood Birgit Rathkolb,1,2 Helmut Fuchs,1 Valérie Gailus-Durner,1 Bernhard Aigner,2 Eckhard Wolf,2 and Martin Hrabˇe de Angelis1,3,4 1

Institute of Experimental Genetics, German Mouse Clinic, Helmholtz-Zentrum München, German Research Center for Environmental Health, GmbH, Neuherberg, Germany 2 Institute of Molecular Animal Breeding and Biotechnology, Gene Center, Ludwig-Maximilians-Universität München, Munich, Germany 3 Institute of Experimental Genetics, Life and Food Science Center Weihenstephan, Technische Universität München, Freising-Weihenstephan, Germany 4 German Research Center for Diabetes Research, Neuherberg, Germany

ABSTRACT Basic phenotyping of inbred mouse strains and genetically modified mouse models usually includes the determination of blood-based parameters as a diagnostic screen for genotype effects on metabolism and organ function. A broad range of analytes, including hematological parameters, can be reliably determined in mouse blood, if appropriate samples are available. Here we describe recommended techniques for blood collection from mice and the considerations that have to be taken into account to get adequate samples for hematological investigations. Furthermore, we describe established methods used in the German Mouse Clinic (GMC) to determine hematological parameters in the C 2013 by John Wiley & Sons, Inc. mouse. Curr. Protoc. Mouse Biol. 3:101-119 Keywords: mouse r blood sample collection r hematology

INTRODUCTION The determination of diagnostic values in mouse blood plays a key role in mouse phenotyping (Justice, 2008; Gailus-Durner et al., 2009; Wakana et al., 2009). Besides hematological data, which are used to discover alterations of hematopoiesis or blood cell morphology in mice, a broad range of plasma parameters can be measured in a single blood sample, if accurately planned. However, limitations occur due to the relatively small sample volume that can be collected from a mouse, and by the fact that different measurements require distinct procedures of sample processing. Here, we first describe our recommended method of blood collection from mice by puncturing the retro-bulbar venous plexus of anesthetized mice. In some countries, this technique might not be allowed in the case of non-terminal blood collection. Therefore, alternative techniques, like bleeding of mice from the submandibular, sublingual, or tail vein, will be briefly described and discussed, without giving a detailed protocol. Furthermore established protocols used in the German Mouse Clinic (GMC; GailusDurner et al., 2005) for the determination of basic hematological parameters in mice are described. Details on serum and plasma preparation are given in our contribution on the analysis of clinical chemistry and other laboratory tests on mouse plasma (Rathkolb et al., 2013). All protocols in this article include information on requirements concerning blood sample processing and sample quality for valid results. Mouse Blood Collection and Analysis Current Protocols in Mouse Biology 3: 101-119, June 2013 Published online June 2013 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/9780470942390.mo130054 C 2013 John Wiley & Sons, Inc. Copyright

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Blood collection from mice (Basic Protocol 1 and Alternate Protocols 1-3) Collecting blood samples from mice demands technical skills that can only be acquired by sufficient practice. Therefore, the availability of experienced personnel for blood collection is necessary. Samples collected by untrained persons often do not match the requirements concerning volume and quality for the analyses planned. The decision on which method is the best to be used for blood collection in a certain experiment depends on a variety of considerations: 1. Which methods are allowed according to relevant animal welfare legislation? 2. What is the required sample volume? 3. Are the mice killed after bleeding or should they stay alive? 4. How do the different methods of anesthesia, blood collection, and sample preparation affect the parameters to be analyzed? In Basic Protocol 1, we describe the puncture of the venous plexus behind the eye ball (retro-bulbar bleeding) as the standard method used in the German Mouse Clinic (GMC). Due to national differences in animal welfare legislation, alternative methods of blood collection might be preferred. In addition, blood collection from conscious mice may be necessary for specific analyses, where the use of anesthesia may prevent the measurement of valid results. Possible alternative methods are blood collection from mice by puncture of the submandibular vein (Alternate Protocol 1), the sublingual vein (Alternate Protocol 2), or the lateral tail vein (Alternate Protocol 3). Methods that can be applied for terminal blood collection in deeply anesthetized animals or directly after euthanasia include heart puncture and the puncture of the caudal vena cava. A detailed description of various methods to collect blood from mice and other laboratory animals is included in Current Protocols in Immunology (Donovan and Brown, 2006); therefore, we will give only a short description here of selected alternative methods.

Automated hematological analysis of mouse blood (Basic Protocol 2 and Alternate Protocol 4) While not explicitly required, modern/automated equipment significantly facilitates thorough, precise, and accurate blood analysis. For example, dilution followed by manual counting (e.g., using a hemacytometer) permits peripheral white and red blood cell counting—however, it is far more efficient to use an appropriate automated hematology analyzer, particularly when handling more than a few samples at a time and/or when more than a single readout is desired. In specific instances, traditional analytical approaches are still recommended, and we discuss this in greater detail in the Commentary.

Mouse Blood Collection and Analysis

In general, the same technologies as used for human blood analysis are applied to mouse blood, e.g., impedance and photometric techniques that might be combined with flow cytometric methods after fluorescence staining of cells. Mouse blood, however, is considerably different from human blood. Erythrocyte size is smaller (about 50 fl in mice versus 80 fl in humans) and red blood cell counts are higher in mice (8–11 × 106 /μl in mice versus 3–5 × 106 /μl in humans). Differences also occur in platelet size (5 to 7 fl in mice versus 7 to 10 fl in humans) and numbers (700–1500 × 103 /μl in mice versus 13—360 × 103 /μl in humans). Therefore, devices should be used that are equipped with special software for mouse blood analysis, ensuring the application of feasible dilutions and the correct discrimination between cell populations. However,

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due to technical effects, values obtained using different devices can significantly differ. Therefore, it is strongly recommended to use the same device for all samples analyzed within one project. Within the Munich ENU Mouse Mutagenesis project, the German Mouse Clinic, and the EUMODIC project (Rathkolb et al., 2000; Gailus-Durner et al., 2009; Gates et al., 2011), we have successfully used the “Scil Vet abc” (Scil animal care company, http://www.scilvet.nl/), the ADVIA 120 and 2120 (Siemens), and the Sysmex XT2000iV (Sysmex, https://www.sysmex.com/) systems to analyze mouse blood samples. The first device mentioned uses only impedance technology and photometry and is useful for determining a basic peripheral blood cell count from only 12 μl of whole blood (Aigner et al., 2011). However, the differentiation of white blood cells based on cell size only does not provide reliable differential leukocyte counts, and no information on reticulocyte numbers is obtained. In contrast, the ADVIA systems and the Sysmex XT2000iV system apply additional flow-cytometric techniques, adding information on differential white blood cell counts and reticulocyte counts (Lilliehook and Tvedten, 2009a,b; Ameri et al., 2011). However, the sample volume needed for these devices is higher: the ADVIA system needs 160 to 175 μl of whole blood, while the Sysmex system needs 85 μl of whole blood. In both systems, the use of prediluted samples is possible. For the Sysmex system, capillary tubes, consisting of sample tubes prefilled with a diluent and 50 μl end-to-end capillaries, are available for the analysis of 1:5 diluted samples in the so-called “capillary mode." In Basic Protocol 2, we describe the use of the Sysmex XT 2000iV system; the use of the “Scil Vet abc” or “Scil Vet abc Plus” system is described in Alternate Protocol 4.

BLOOD COLLECTION FROM MICE BY RETRO-BULBAR PUNCTURE Blood collection from the venous plexus inside the orbit behind the eyeball is currently the most commonly used method to get blood samples from mice in Germany. The advantages of this method are: (1) suffering of the mice due to the procedure when carried out by trained staff is within acceptable limits; (2) adequate amounts of highquality blood can be collected with a minimal incidence of hemolysis; (3) since only short-term anesthesia is required, the collection of samples from up to 20 mice within 1 hr is possible.

BASIC PROTOCOL 1

However, there is some controversy surrounding the technique due to possible side effects of isoflurane anesthesia and the risk of injuries of the eye. In some countries, retro-bulbar bleeding is allowed as a terminal bleeding method only. In this case, an injection narcosis must be applied, or isoflurane narcosis must be applied continuously via a face mask to ensure deep narcosis until the animal is killed by blood withdrawal. Here we describe the protocol applied in the German Mouse Clinic including notes on alternative methods, where appropriate.

Materials Mice Isoflurane Oxygen or compressed air Laboratory scale, convenient for body mass determination of mice (e.g., Kern 440-47N, Precision scale; Kern & Sohn GmbH, http://www.kern-sohn.com ) Isoflurane vaporizer (e.g., Penlon Sigma Delta Vaporizer, UNO Roestvaststaal BV, Zenvenaar, Netherlands) Flowmeter for oxygen (O2 ) and nitric oxide (N2 O) and/or compressed air (e.g., Flowmeter Type SF1, SF2, or SF3; UNO Roestvaststaal BV, http://www.unobv.com/)


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Anesthesia induction chamber for mice (e.g., UNO Roestvaststaal BV, http://www.unobv.com) Small glass capillaries (about 1 mm in diameter and 20 to 30 mm long; e.g., NeoLab Migge GmbH, http://www.neolab.de/) 0.5- to 1.0-ml sample tubes (e.g., Kabe Labortechnik GmbH, http://www.kabe-labortechnik.de/): depending on the analyses planned plain tubes or heparin coated and/or EDTA coated tubes must be used (see protocols for sample analyses) Anesthetize mouse Refer to Figure 1. 1. Determine body mass of mice to calculate sample volume maximum. Body mass determination is not necessarily needed if terminal blood collection is planned, since in this case blood withdrawal is continued until the mouse is dead.

2. Open the gas supply and set air flow to approximately 1.5 liters/min (compressed air), set vaporizer to add 5% isoflurane to the gas flow for narcosis induction, place mouse in the anesthesia induction chamber, and wait ∼2 to 3 min until it is deeply anesthetized. A sufficiently deep anesthesia is present when the mouse has stopped moving, skeletal muscles are relaxed and a decreased frequency but increased depth of breathing is visible. For terminal blood collection, a longer-lasting anesthesia is needed. This is achieved by an injectable anesthetic e.g., ketamine/xylazine, or by continuously applying the isoflurane anesthesia using a face mask.

Figure 1 Equipment for short-term anesthesia of mice using isoflurane. The isoflurane evaporator is connected to a supply of compressed air via the flowmeter. Gas flow is set to 1.3 to 1.5 l/min and 5% (v/v) isoflurane is added by the evaporator. For anesthesia, mice are placed in the induction chamber for 2 to 3 min.


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Collect blood sample 3. Remove the mouse from the induction chamber. Fix the skin of the neck between the index finger and thumb of the left hand (right hand for left-handed workers), thereby carefully tightening the skin around the throat to achieve a mild compression of the jugular vein. You might additionally fix the tail between the ring finger and the little finger.

4. Insert a glass capillary into the medial corner of the eye (preferable the right eye for right-handed persons) in the space between the bulbus and the eyelid, without touching the cornea. 5. Push the capillary gently forward in the direction of the base of the opposite ear by rotating it simultaneously between the index finger and thumb of the right hand (left hand for left-handed workers), until blood enters the capillary. 6. Collect the drops of blood in prepared sample tube(s) until the desired volume is reached. Since the anesthesia lasts only for 45 to 60 sec after removal of the mouse from the induction chamber, personnel have to be adequately trained to be able to collect the required volume within this time span. Sample tubes have to be chosen according to the requirements of the tests to be performed. Li-heparin-coated or EDTA-coated sample tubes are provided by several suppliers.

7. Loosen the grip on the skin in the neck of the mouse to relieve the vein compression. Remove the capillary from the eye and apply a gentle pressure to the closed eye of the mouse using a clean tissue to stop bleeding before putting the mouse back to its cage. In case of a non-terminal bleeding, mice should be observed after the bleeding procedure until they reach complete consciousness and checked for well-being before being transported back to the mouse room.

8. Process samples as recommended for the measurements planned (see protocols for sample analyses) Sample processing depends on the investigations planned. For hematological investigations, EDTA-coated sample tubes are used for blood collection. EDTA-treated blood samples for hematological analyses are kept in motion at room temperature using a rotary agitator. For parameters measured in serum or plasma, either plain sample tubes or EDTA- or Li-heparin-coated tubes and adequate sample processing are required. Details are given in our contribution on the analysis of clinical chemistry and other laboratory tests on mouse plasma or serum (Rathkolb et al., 2013).

BLOOD COLLECTION FROM MICE BY PUNCTURE OF THE SUBMANDIBULAR VEIN

ALTERNATE PROTOCOL 1

Blood collection from the submandibular vein was reported as a suitable alternative method to retro-bulbar puncture for collecting larger blood samples from anesthetized or conscious mice (Golde et al., 2005). However, several studies indicated problems that can occur with this technique, including difficulties in hitting the correct point of puncture in the first attempt, especially in conscious mice, which might be struggling heavily, as well as excessive bleeding and subcutaneous hemorrhages, mainly due to accidental puncture of the accompanying artery, especially in mice with impaired blood clotting (Heimann et al., 2010; Holmberg et al., 2011). The latter event also leads to the collection of a mix of arterial and venous blood. Mouse Blood Collection and Analysis


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Additional Materials (also see Basic Protocol 1) Sterile 4- to 5-mm lancets (available, e.g., from B. Braun Melsungen AG, http://www.bbraun.com/) NOTE: While this method can be performed in conscious mice, we strongly recommend anesthesia both to minimize trauma to the animal and to facilitate collection by the researcher. 1. Grasp the animal at the tail and also grip the skin on the back and neck to trigger venous congestion and immobilization of the animal’s head and body. Conscious mice often need rather firm handling to prevent excessive struggling; we strongly recommend using an anesthetic compatible with experimental procedures.

2. Once the animal is in position, use a sterile lancet to puncture the caudal contour of the mandible approximately 3 mm caudal and 1 mm dorsal to the lateral tactile hair. 3. Collect drops of blood in the sample tube(s). 4. Once collection is complete, stop the bleeding by releasing the grip on the animal and applying gentle pressure to the puncture site. ALTERNATE PROTOCOL 2

BLOOD COLLECTION FROM MICE BY PUNCTURE OF THE SUBLINGUAL VEIN This method, first used in rats, is a recommended alternative method to retro-bulbar puncture in mice, and has to be carried out in anesthetized mice (Heimann et al., 2010; Seibel et al., 2010). It is suitable for collecting large blood volumes, and usually has only a minor impact on the animal‘s well-being. However, using this technique in mice successfully requires some practice, as is the case with the retro-bulbar puncture method (Basic Protocol 1).

Additional Materials (also see Basic Protocol 1) Cotton swabs 24-G hypodermic needles Additional reagents and equipment for ketamine/xylazine anesthesia of mice (Donovan and Brown, 1998) NOTE: For this blood collection method, we strongly recommend having two people working in tandem. 1. Anesthetize the mouse by ketamine/xylazine injection (Donovan and Brown, 1998). 2. Once mouse is anesthetized, have one person grasp the mouse from the back and hold the animal down in a supine (belly-up) position. 3. While the first person restrains the mouse in a supine position, have the second person open the animal’s mouth and pull the tongue out; the second person should maintain a grip on the tongue using his or her thumb and a cotton swab. 4. While holding the tongue out as described in the previous step, have the second person puncture the thick caudal part of the sublingual vein with a 24-G needle. 5. To avoid complications such as swelling of the tongue and hematomas at the puncture site, avoid puncturing the sublingual vein too close to the tongue’s apex. Mouse Blood Collection and Analysis

6. Have the first person then turn the animal over in a ventral position while the second person immediately starts collecting the drops of blood in a sample tube.

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BLOOD COLLECTION FROM MICE BY INCISION OF THE LATERAL TAIL VEIN


Blood collection from the lateral tail vein is the only method recommended for blood collection from conscious mice (Aasland et al., 2010; Christensen et al., 2009). A detailed description of this method has been published in Current Protocols in Molecular Biology (Argmann and Auwerx, 2006). With this method, only small samples (50 to 200 μl) can be easily collected. It is especially useful for the repeated collection of small samples from the same mouse, e.g., in a glucose tolerance test.

Additional Materials (also see Basic Protocol 1) Mouse restraining device Scalpel with sterile scalpel blade Hematocrit capillaries or 0.5- to 1.0-ml sample tubes 1. Fix mouse in a restraining device, ensuring that the tail remains accessible for blood collection. Immersing the tail in warm water prior to incision facilitates blood flow and thus collection. For longer-term/repeated sampling, use dry/non-contact methods to heat the tail (if necessary), avoiding direct contact with the incision site.

2. Using a sharp, sterile scalpel, make a small lateral incision at the cranial part of the tail. 3. Collect droplets of blood with hematocrit capillaries or sample tubes.

HEMATOLOGICAL ANALYSIS OF MOUSE BLOOD USING THE SYSMEX XT 2000iV SYSTEM

BASIC PROTOCOL 2

The Sysmex XT 2000iV system (Fig. 2) is an automated blood cell counter for veterinary use. A variety of predefined settings for different animal species, including mouse and rat, can be chosen from the menu of the control software. Additionally, custom-made settings can be set and stored if the predefined settings do not fit the specific characteristics of the samples analyzed, e.g., in case of abnormal cells occurring in blood. The analyzer offers three different modes of sample input: the automated mode, the manual mode, and the capillary mode. The automated mode is intended to be used for the automated analysis of high numbers of samples, which are automatically mixed before the sample for the measurement is drawn. This mode requires the use of 5-ml sample tubes and a minimum volume of 200 μl of whole blood. Therefore, it is not feasible to use this mode with mouse samples. Using the manual mode, 85 μl of whole blood is drawn from the sample tube, which has to be manually placed below the sampling needle for aspiration. In the capillary mode, handling is the same as with the manual mode, but the software takes the dilution of the sample into account and calculates the values of undiluted whole blood from the measurement of diluted samples. This mode has to be used with 1:5 diluted samples only.

Materials Mice Quality-control blood samples (Sysmex E-Check high level, medium level and low level, delivered with target values and limits on CD) Adapted reagents packs for the Sysmex analyzer (Sysmex):



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Required for complete blood counts and differential white blood cell counts: Cellpack (cat. no. 83400116) Stromatolyser FB (cat. no. 94404613) Stromatolyser 4DS (cat. no. 98417216) Stromatolyser 4DL (cat. no. 98417615) Sulfolyser (cat. no. 90414114) For additional reticulocyte counts: Retsearch II (cat. no. 98416211) Cell clean solution (Sysmex cat. no. 83401621; https://www.sysmex.com/) EDTA-coated sample tubes (e.g., Kabe Labortechnik, http://www.kabe-labortechnik.de/, EDTA 500 A or EDTA 1000 A Standrand) or capillary tubes (prefilled with Sysmex Cellpack buffer as diluent) and 50-μl end-to-end capillaries (Sysmex Deutschland GmbH, cat. no. 99940020, https://www.sysmex.com/) Overhead rotary agitator for small sample tubes (e.g., Multi RS-60; BioSan, http://www.biosan.lv/) Sysmex XT 2000iV analyzer (Sysmex, https://www.sysmex.com/) Additional reagents and equipment for blood collection (see Basic Protocol 1 and Alternate Protocols 1-3)


Figure 2 Equipment used in the German Mouse Clinic for hematological analyses of mouse blood. (A) The Sysmex XT 2000iV system for analyses of animal blood samples. (B) Using the manual mode, a blood sample is placed below the sampling needle for sample aspiration, which is started by pressing the blue button behind the sampling needle. (C) Screenshot of the main menu of the control software: for QC-sample analysis open QC analysis menu (down right), to start sample analysis, choose manual menu (headline, second symbol).

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1. Collect 100 μl of blood from each mouse into EDTA-coated sample tubes, or fill an end-to-end capillary with 50 μl blood and put it into a sample tube with dilution medium (capillary tubes for Sysmex capillary mode). The same method of blood collection (see Basic Protocol 1 and Alternate Protocols 1-3) should be used for all samples tested. Methods involving intense tissue contact of blood can impair the measurement, due to the formation of coagula, and severe hemolysis will also alter the blood cell counts. Since the degree of hemolysis cannot easily be determined by eye in whole-blood samples, methods of blood collection should be selected that have been used previously for the successful preparation of hemolysis-free plasma sample.

2. Mix samples gently by inverting them several times and place them on a rotary agitator at room temperature to keep them in motion. 3. Start the Sysmex XT 2000iV system and log in to the control software. Wait until the working temperature is reached, then make sure that the device is working properly by analyzing the three control blood samples (Fig. 2A and B). Check that all reagent supplies are sufficiently filled and appropriately connected to the analyzer. Missing connections or empty supplies will give an alarm note on the monitor during the starting procedure. Before the control blood samples can be analyzed, the lot number of the samples, as well as target values and limits, have to be transferred from a disc delivered with the samples to the control software as described in the manual. For the analysis of control samples, select QC analysis from the main menu (Fig. 2C), choose the correct lot number, and measure the sample. Values deviating from the target range will be highlighted in yellow (borderline level) or red (level outside the limits). If the control analysis leads to correct values (Fig. 3A), accept results; otherwise, check for possible causes of failure, e.g., expired reagents or controls or technical problems.

4. For sample analysis, choose the “manual” button and enter the number of the first sample to be analyzed in “sample number.” Choose the correct settings for the analysis of mouse blood (“animal species”: mouse) and the measurement mode of choice (“mode”: manual or capillary, respectively) from the manual menu of the control program. Options can also be chosen as to whether only the complete blood count (CBC) should be measured or reticulocyte counts (Ret) and/or differential white blood cell counts (Diff) should be included in the analysis.

5. Analyze samples and check the results for reliability one after the other. Samples are aspirated from the sample tube placed below the sampling needle when the “start button” behind the needle is pressed. Results are stored automatically by the control software and can be displayed for each sample in numbers and plots, using the data browser in the menu (Fig. 3B). Before pressing the “start button,” make sure that the sampling needle is dipping well into the blood to avoid aspiration of air into the system. Starting from the entered number for the first sample, the program automatically creates running sample numbers. Different information, e.g., mouse ID numbers, have to be entered manually for every sample. Results can be stored as an Excel- or database-compatible .csv file and transferred into a database and/or used with software for statistical analyses. The data explorer is used to select results from the archive for external storage or printing.

6. When sample analyses are finished, run the shut-down procedure using cell clean solution to clean the tubes of the analyzer before switching off the system. Mouse Blood Collection and Analysis


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Figure 3 Screenshots of the Sysmex XT 2000iV control software. (A) Results obtained from the analysis of a control blood sample; values that are close to the borders of or not within the acceptable target range would be highlighted in yellow or red, respectively. (B) Results obtained from a mouse blood sample analysis including reticulocyte counts and the differential white blood cell counts.



HEMATOLOGICAL ANALYSIS OF MOUSE BLOOD USING THE “Scil Vet abc” OR “Scil Vet abc Plus” ANIMAL BLOOD COUNTER For the “Scil Vet abc-animal blood counter” (Scil Aminal Care Company) there are two different settings for the analysis of mouse blood available, which are encoded on chip-cards and give remarkably different results. One card is the “laboratory mouse card"; the other is the “C57BL/6 mouse card.” The settings for the analyses are entered by introducing the corresponding chip-card into the chip card reader and choosing the encoded settings via the menu. For the analysis of control blood samples, the chipcard “Blood Control” is used, while for the analysis of mouse-blood samples, one of the two available mouse cards has to be chosen. Results are shown on the display and automatically sent to a printer connected to the device. The analyzer can also be connected

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by a serial interface to a computer, where specific software provided by Scil (Scil-VIP software for Windows 7) or compatible LIMS software is installed for online transfer of data. The device “Scil Vet abc Plus” is delivered with preinstalled settings for different animal species, which are selected via the menu on the touch-screen display of the analyzer. Results are shown on the display, and results from up to 1000 samples are stored within the control software. They can additionally be sent automatically to a connected printer and to LIMS software on a connected computer, as described above. Sample collection and storage are as described in Basic Protocol 2. Since the required sample volume is much smaller for these devices (50 μl of whole blood are sufficient), it is not recommended to use diluted samples. However, if diluted samples are used with the “Scil Vet abc-animal blood counter,” whole blood values have to be calculated manually from the results obtained for diluted samples. An additional measurement of diluted samples is advisable for the determination of more exact platelet counts if platelet numbers exceed the range of reliable measurement.

Materials Mice Quality control blood samples (Minotrol 16, Control H–high level–and Control L–low level; Scil Animal Care Company, http://www.scilvet.nl) EDTA-coated sample tubes (e.g., from Kabe Labortechnik GmbH, http://www.kabe-labortechnik.de/, EDTA 500 A or EDTA 1000 A Standrand) Over-head rotary agitator for small sample tubes, e.g., “Multi RS-60” (BioSan, http://www.biosan.lv/) “Scil Vet abc” or “Scil Vet abc Plus” (Scil Animal Care Company, http://www.scilvet.nl), including an Abc-reagent-pack, which contains all necessary reagents, and a refuse bag for the waste liquids Minoclair solution (Scil Animal Care Company, http://www.scilvet.nl) Additional reagents and equipment for blood collection (see Basic Protocol 1 and Alternate Protocols 1-3) 1. Collect 50 μl of mouse blood into EDTA-coated sample tubes as described above, mix thoroughly, and place them on the rotary agitator. 2. Start the analyzer and make sure that it is working properly by analyzing the control blood samples. For the analysis of control blood samples using the “Scil Vet abc” system, the specific settings for control blood are loaded from the “Blood Control” chip-card. Using the “Scil Vet abc Plus” system, the settings are selected via the menu.

3. Switch settings for the analysis of mouse blood samples by introducing the appropriate chip-card or choosing this animal species from the menu 4. Analyze mouse blood samples and check the results obtained for reliability. Sample numbers are automatically counted up; additional information, like mouse ID, can be entered via the integrated keyboard.

5. When measurements are finished, run the special cleaning procedure, selectable via the menu, using Minoclair solution to clean the conduction tubes of the analyzer, before switching off the device. Mouse Blood Collection and Analysis


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COMMENTARY Background Information Background information for blood collection High-quality and adequately processed blood samples are an important precondition for reliable results of blood analyses. Since measured values can be altered by a variety of parameters during the whole process of sample collection and processing, e.g., environmental stress, anesthesia, and methods of sample collection and sample preparation, it is essential to use a standardized protocol for all animals within a study (Nemzek et al., 2001; Fernández et al., 2010). According to the recommendations of the German Society of Laboratory Animal Science (GV-SOLAS), the sample volume should not exceed 10% of the whole blood volume, if the animals are to stay alive. The total blood volume of mice is ∼70 μl to 80 μl per g body weight, resulting in an estimated total blood volume of 1.7 to 2.0 ml for a mouse weighing 25 g; this means a maximal sample volume of 200 μl. In exceptional cases, up to 20% of the total blood volume (400 μl for a mouse of 25 g) can be collected with sufficient fluid substitution (prewarmed, sterile isotonic saline i.p.). If a repeated collection of small blood samples is planned, catheterization or even automated sampling methods might be preferred (Xie et al., 2003). In the case of terminal blood collection, mice are deeply anesthetized, preferably by injectable anesthetics, and killed by blood withdrawal. Using this option, more than 1 ml of blood can be collected from a single mouse.


Background information for automated analysis of mouse blood Alterations of hematological parameters can result from primary effects on hematological cell function impacting cell division, differentiation, or life span, or occur secondarily due to iron metabolism disorders, kidney diseases, acute or chronic bleeding, infection, or malnutrition. The inter-individual variation of age-matched mice belonging to the same inbred strain and housed under identical conditions is low for many hematological parameters. Therefore, quite small effects can be detected reliably in well organized experiments. During the second half of the 20th century, automated blood cell counts replaced the older manual analysis of hemoglobin concentrations and microscopic techniques, which were based

mainly on the determination of cell counts in diluted or hemolyzed samples using counting chambers and on the light-microscopic evaluation of specifically stained blood smears (Nelson and Lamont, 1961; Nelson and Carville, 1962; Rappaport et al., 1988). Hematology analyzers for veterinary use became available in the last two decades of the 20th century (Knoll and Rowell, 1996), with several manufacturers providing specific settings for the analysis of mouse blood. While automated determination of the red blood cell count and size results in an increased accuracy compared to manual methods, the reliability of automated white blood cell differentiation was controversial (Simmons et al., 1974; Goossens et al., 1991; Hyun et al., 1991). Especially for the diagnosis of characteristic morphological features of blood cells indicating certain diseases, the microscopic evaluation of blood smears is still the method of choice (Allison and Meinkoth, 2007).

Critical Parameters and Troubleshooting Important considerations for blood collection All methods of blood collection and processing may influence the outcome of analyses conducted on these samples. Some parameters will give invalid values if the animal was stressed before or during blood collection, or if the sample is hemolytic (Fig. 4). Blood sampling methods involving the direct contact of blood with the skin, and underlying tissues will activate the blood clotting system and result more frequently in coagulation and hemolyzed samples. Also, the method chosen for anesthesia might influence the outcome of blood testing. Isoflurane anesthesia, for example, has been shown to cause impaired insulin secretion and glucose tolerance (Tanaka et al., 2009), and ketamine also seems to have effects on glucose metabolism (Shin et al., 2006). Methods that are associated with muscle injuries (blood vessel puncture in situ, heart puncture, submandibular vein puncture) can result in altered values of several parameters (e.g., muscle enzyme activities) measured in the sample. The special conditions required for the reliable determination of specific parameters are described, where necessary, in the respective protocols for blood sample analyses. Equipment used in the GMC for sample collection and agitation is shown in Figure 5.

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Figure 4

Plasma samples showing mild (right) to moderate (left) hemolysis.

Important considerations for automated analysis of mouse blood Hematological values depend on the genetic background of the mice tested (WirthDzieciołowska ˛ et al., 2009), the sampling site (Nemzek et al., 2001), sample preparation, and the device used for the analysis. Therefore, it is necessary to apply a defined standard operating procedure (SOP) for all animals investigated within a study and to include suitable controls in all experiments. To prevent the formation of micro-coagula, EDTA-treated blood should be used. After blood collection into EDTA-coated sample tubes and/or dilution with EDTA-containing saline, samples are mixed thoroughly to achieve a homogenous distribution of the anticoagulant. The samples are kept in motion at room temperature using a rotary agitator until measured. Undiluted blood samples should be analyzed within 2 to 3 hr after collection, while diluted samples can be stored for up to 24 hr. Severe hemolysis in the samples investigated will falsify the results of the red blood cell count and erythrocyte indices. The formation of micro-coagula can change the red blood cell, platelet, and leukocyte counts, and creates a risk of internal tubes of the analyzer becoming obstructed. This becomes obvious from missing values in the result set. If automated

flushing of the system does not remove the clot, the system has to be thoroughly cleaned by the instrument-specific cleaning procedure before further samples are analyzed. If the sample volume provided is too small or if the sampling needle does not dip properly into the blood during sample aspiration, the amount of blood aspirated might not be sufficient, leading to an inadequate dilution of the sample during the measurement. This results in unusually low results for all cell counts and hemoglobin and hematocrit values. In these cases, it is recommended to ensure correctness of results by a second measurement from the same sample or an additional sample collected from the same mouse.

Anticipated Results Anticipated results for blood collection Generally speaking, any of the manual procedures outlined in this manuscript (Basic Protocol 1 and/or Alternate Protocols 1-3) should result in collection of sufficient quantities of high-quality (hemolysis-free) blood suitable for downstream analysis. Anticipated results for automated analysis of mouse blood The results obtained by the analyses described above are data of the standard complete



Volume 3

Figure 5 Equipment used in the German Mouse Clinic for mouse blood collection. (A) Li-heparincoated (orange cap) and EDTA-coated (red cap) sample tubes used for the collection of samples for clinical-chemical and hematological analyses, respectively. (B) Overhead rotary mixer “Multi RS-60” (BioSan, http://www.biosan.lv/), used to keep blood samples for hematological analyses in motion until analyzed.

blood count, consisting of the following parameters:


Total white blood cell count (WBC) Platelet count (PLT) Mean platelet volume (MPV) Total red blood cell count (RBC) Mean corpuscular volume (MCV), which approximates the mean volume of erythrocytes Mean cellular hemoglobin content of erythrocytes (MCH)

Mean cellular hemoglobin concentration in erythrocytes (MCHC) Red cell distribution width (RDW), a measure of red cell size variation. Additional parameters that are reliably determined by the Sysmex XT 2000iV system are parameters concerning the reticulocyte count (given as total number and percentage of RBC as well as distribution in different maturity stages), the platelets [volume percent of plateletcrit (PCR), platelet distribution width (PDW), and platelet–large cell ratio (P-LCR),

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Normal or increased

Reduced

Increased

Increased

Increased

Increased

Normal

Microcytic

Macrocytic

Polyglobulia

Relative

Secondary

Primary

Osmotic effects

Reduced

Reduced

Slightly reduced

Increased

Normal

Reduced or increased

Possible causes

Increased or normal

Decreased Unusual concentrations of osmotically active or normal substances in blood plasma or blood cells influence intracellular versus extracellular water balance, resulting in cell swelling or shrinkage

Normal or Erythrocyte overproduction due to genetic increased changes in hematopoietic stem cells. Erythropoetin levels are normal or low.

Normal or Secondary to increased erythropoietin synthesis increased due to decreased blood oxygen saturation, specific kidney diseases, or pathologic hemoglobin variants

Relative polycytemia is due to a reduced plasma volume e.g., caused by inadequate fluid uptake and/or polyuria (hemoconcentration)

Impaired cell division during hematopoiesis, e.g., due to folic acid or vitamin B12 deficiency or inborn errors of DNA synthesis

Iron deficiency, impaired globin synthesis (thalassemia), impaired heme synthesis (sideroblastic anemia), spherocytosis

Normal or Intravascular hemolysis, chronic or acute increased bleeding, hyporegenerative anemia, secondary to hemodilution

RDW

Normal or slightly Normal increased

Reduced or normal

Normal or reduced

Normal

MCHC

Normal or Normal or reduced reduced

Normal

Normal

Increased or normal

Reduced

Increased Increased Normal or reduced or reduced

Increased

Increased

Increased

Increased

Reduced

MCH

Normal or Normal slightly increased

MCV

Normal or Reduced reduced

Reduced

Reduced or normal

HCT

a Abbreviations: RBC, red blood cell count; HGB, hemoglobin; HCT, hematocrit; MCV, mean corpuscular volume; MCH, mean corpuscular hemoglobin content; MCHC, mean corpuscular hemoglobin concentration; RDW, red cell distribution width.

Normal

Increased

Increased

Increased

Increased

Reduced

Reduced

Reduced

Reduced

Erythropenic, normocytic

HGB Reduced

RBC

Anemias

Findings

Table 1 Possible Findings Concerning the Red Blood Cell Counta

giving the percentage of platelets >12 fl], and the differential white blood cell counts consisting of total numbers and proportions of WBC in percent for the following cell types: Lymphocytes (Lymph) Monocytes (Mono) Neutrophil granulocytes (Neut) Eosinophil granulocytes (Eo) Basophil granulocytes (Baso). Since the inter-individual variation of many hematological parameters is quite small in mice belonging to the same inbred strain, even small changes can be detected in a well planned experiment (Tsuchiya et al., 2004; del Barco Barrantes et al., 2006). The analysis of hematological parameters was successfully used in the Munich ENU mouse mutagenesis project to establish new mutant mouse lines with aberrant phenotypes (Hrab˘e de Angelis et al., 2000; Aigner et al., 2011). A broad spectrum of hematological changes can be diagnosed. Possible findings concerning the red blood cells are listed in Table 1. Leukocyte counts can be increased (leukocytosis, leukosis) or reduced (leukopenia). Simultaneously, the composition of leukocytes in the peripheral blood can be affected, resulting in changes in the differential white blood cell count. Similarly, platelet numbers can be increased (thrombocytosis) or decreased (thrombocytopenia), and/or platelet morphology can be changed (decreased or increased MPV values and changes in platelet size distribution). However, hematological parameters can change due to a wide variety of different causes. Genetic alterations might directly affect proliferation and/or differentiation of hematological stem cells, e.g., due to altered responsiveness to or signaling of growth factors (Divoky et al., 2001; Ruan et al., 2005) or impaired cell division (Devlin et al., 2010). Hematological changes can also occur due to altered morphology, life-span, and/or function of blood cells (Muro et al., 2000; Shet et al., 2008), impaired hemoglobin synthesis (Bishop et al., 2011) secondary to changes in regulatory pathways (Haase, 2006) or iron metabolism (Mok et al., 2006; Vujić Spasić et al., 2007), primary diseases of other organs (Fernandez-Banares et al., 2009; GonzalezCasas et al., 2009), or infections (Rivera and Ganz, 2009). Therefore, additional investigations, like clinical chemistry analyses and/or

microscopic evaluation of blood cells (blood smears or cytospin preparations), and/or studies of cellular properties in culture, are usually necessary to elucidate the cause of changes detected in the peripheral blood cell count.

Time Considerations Time considerations for blood collection Using the protocols described above, it is possible for one or two technicians to collect blood samples of sufficient quality from an experimental group of about 40 mice within 2 to 3 hr. By retro-bulbar bleeding, a trained technician is able collect sufficient amounts of blood of high quality (hemolysis-free) from one mouse within 2 to 4 min. Therefore, about 20 samples can be collected within 1 hr using this method. If injection anesthesia is carried out, additional time for the injection of the anesthetics and the time until the animals are sufficiently anesthetized has to be considered. In this case, it is necessary to treat the eyes of the anesthetized animals after blood collection using a suitable ointment if the mice are not to be killed. If the samples have to be processed or analyzed within a certain time period after collection, additional personnel should proceed with sample processing and analyses in parallel, if large groups of animals are investigated within 1 day. Time considerations for automated analysis of mouse blood The time to analyze a single sample depends on the device used. The Scil Vet abc system needs 2 min for the analysis of one sample. Therefore, up to 30 samples can be analyzed within 1 hr. The Sysmex XT 2000iV system needs less than a minute for the analysis of one sample, increasing the throughput to up to 80 samples per hour.

Acknowledgments We thank Kateryna Micklich and Elfi Holupirek for supporting the measurements and technical advice for the manuscript. This work has been funded by the German Federal Ministry of Education and Research to the German Center for Diabetes Research (DZD e.V.) and to the GMC [NGFNplus grant No. 01GS0850, 01GS0851 and Infrafrontier grant 01KX1012)] as well as by an EU grant (EUMODIC, LSHG-2006-037188, German Mouse Clinic).


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Holmberg, H., Kiersgaard, M.K., Mikkelsen, L.F., and Tranholm, M. 2011. Impact of blood sampling technique on blood quality and animal welfare in haemophilic mice. Lab. Anim. 45:114120.

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Wirth-Dzieciołowska, ˛ E., Py´sniak, K., Smolińska, M., and Gajewska, M. 2009. Selected peripheral blood cell parameters in twelve inbred strains of laboratory mice. Anim. Sci. Pap. Reports 27:6977. Xie, F., Bruntlett, C.S., Zhu, Y., Kissinger, C.B., and Kissinger, P.T. 2003. Good preclinical bioanalytical chemistry requires proper sampling from laboratory animals: Automation of blood and microdialysis sampling improves the productivity of LC/MSMS. Anal. Sci. 19:479485.



Volume 3

Hematological assessment in pet guinea pigs (Cavia porcellus): blood sample collection and blood cell identification.

Interferences from blood collection tube components on clinical chemistry assays.

Hazards of blood collection.

Blood collection techniques.

CTC-mRNA (AR-V7) Analysis from Blood Samples-Impact of Blood Collection Tube and Storage Time.

Sterility of blood collection tubes.

Data on the impact of the blood sample collection methods on blood protein profiling studies.

Repeated Blood Collection for Blood Tests in Adult Zebrafish.

Laboratory Diagnostics and Quality of Blood Collection.

Collection and use of autologous fetal blood.

Impact of blood collection and processing on peripheral blood gene expression profiling in type 1 diabetes.

Spuriously low plasma propranolol concentrations resulting from blood collection methods.

Sources of preanalytical error in pharmacokinetic analyses - focus on intravenous drug administration and collection of blood samples.

Influence of blood collection systems on coagulation tests.

On the improvement of blood sample collection at clinical laboratories.

Relevance of EDTA carryover during blood collection.

[Geomatics in the service of blood collection].

Evacuated blood-collection tubes--the backflow hazard.

Blood specimen collection tubes for coagulation tests.

Blood specimen collection tubes for coagulation tests.

Blood specimen collection tubes for coagulation tests.

Comparison of Submental Blood Collection with the Retroorbital and Submandibular Methods in Mice (Mus musculus).

Chemical and hematological changes in stored CPD blood.

Blood gas and hematological changes in experimental peracute porcine pleuropneumonia.