thoughts and progress - nci hub...index of hemolysis (mih nozzle) values were 0.292 ± 0.249, 0.021...

Thoughts and Progress

Multilaboratory Study of Flow-InducedHemolysis Using the FDA Benchmark

Nozzle Model

*Luke H. Herbertson, †Salim E. Olia,†Amanda Daly, ‡Christopher P. Noatch,

‡§William A. Smith, †¶Marina V. Kameneva,and *Richard A. Malinauskas

*Center for Devices and Radiological Health, USFood and Drug Administration, Silver Spring, MD;

Departments of †Bioengineering and ¶Surgery,University of Pittsburgh, Pittsburgh, PA; ‡Lerner

Research Institute, Cleveland Clinic; and§Engineering Department, Perfusion Solutions, Inc.,

Cleveland, OH, USA

Abstract: Multilaboratory in vitro blood damage testingwas performed on a simple nozzle model to determine howdifferent flow parameters and blood properties affectdevice-induced hemolysis and to generate data for com-parison with computational fluid dynamics-based predic-tions of blood damage as part of an FDA initiative forassessing medical device safety. Three independent labora-tories evaluated hemolysis as a function of nozzle entrancegeometry, flow rate, and blood properties. Bovine bloodanticoagulated with acid citrate dextrose solution (2–80 hpost-draw) was recirculated through nozzle-containing andpaired nozzle-free control loops for 2 h. Controlled param-eters included hematocrit (36 ± 1.5%), temperature(25°C), blood volume, flow rate, and pressure. Threenozzle test conditions were evaluated (n = 26–36 trialseach): (i) sudden contraction at the entrance with a bloodflow rate of 5 L/min, (ii) gradual cone at the entrance witha 6-L/min blood flow rate, and (iii) sudden-contractioninlet at 6 L/min. The blood damage caused only by thenozzle model was calculated by subtracting the hemolysisgenerated by the paired control loop test. Despite highintralaboratory variability, significant differences amongthe three test conditions were observed, with the sharpnozzle entrance causing the most hemolysis. Modifiedindex of hemolysis (MIHnozzle) values were 0.292 ± 0.249,0.021 ± 0.128, and 1.239 ± 0.667 for conditions i–iii,

respectively. Porcine blood generated hemolysis resultssimilar to those obtained with bovine blood. Although theinterlaboratory hemolysis results are only applicable forthe specific blood parameters and nozzle model used here,these empirical data may help to advance computationalfluid dynamics models for predicting blood damage. KeyWords: Hemolysis—Red blood cell damage—Nozzlemodel—Fluid dynamics—In vitro hemolysis testing—Computational fluid dynamics.

Many blood-contacting medical devices, includingprosthetic heart valves, catheters, blood pumps, andcardiopulmonary bypass oxygenators, can causeblood cell damage (1–4). Hemolysis, which is charac-terized by the release of intracellular hemoglobinfrom damaged red blood cells (RBCs) into the sur-rounding plasma, can lead to serious clinical sequelaesuch as anemia, hypertension, smooth muscledystonia, arrhythmias, thrombosis, and renal failure(5). To address these potential adverse effects, manu-facturers must demonstrate that their cardiovasculardevices do not cause excessive blood damage beforethey can be marketed. Typically, device safety interms of blood damage potential is demonstratedthrough dynamic in vitro hemolysis testing. Thisusually involves recirculating animal blood through asubject device for a defined duration under clinicallyrelevant flow and pressure conditions and measuringthe amount of resulting hemolysis in relation to thatcaused by a similarly tested, already marketed device(6,7). As many factors, including blood species,temperature, and preparation (e.g., anticoagulation,storage, and blood age), can affect generatedhemolysis (8), it is difficult to compare in vitro testresults from different laboratories and extrapolatethem to clinical outcomes.

While the use of computational fluid dynamics(CFD) modeling is now commonplace for devicedesign, adoption of predictive flow-related blooddamage models requires extensive experimentalvalidation. Empirical studies using various flowmodels (e.g., orifice plate creating a submerged jet,cone-and-plate viscometer, Couette rheometer, cap-illary tube) have shown that flow-related blooddamage is a function of both shear stress and time ofexposure to fluid forces (9–13). Constitutive equa-tions generated by these studies have been incorpo-rated into computational models to predict hemolysis(14) and platelet damage (15) and have been utilized

doi:10.1111/aor.12368

Received January 2014; revised May 2014.Address correspondence and reprint requests to Dr. Luke

Herbertson, US Food and Drug Administration—Center forDevices and Radiological Health, 10903 New Hampshire Ave.,Office of Science and Engineering Laboratories, WO 62, Room2120 Silver Spring, MD 20993, USA. E-mail: [email protected]

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Copyright © 2014 International Center for Artificial Organs and Transplantation and Wiley Periodicals, Inc.

Artificial Organs 2015, 39(3):237–259

mailto:[email protected]


in the development of a variety of medical devices(16,17). However, CFD predictions of blood damageare heavily dependent on which empirical shear-stress exposure time model is used and how it isimplemented (17,18).

CFD analyses are not required in devicepremarketing applications to the US Food andDrug Administration (FDA), but they are increas-ingly being used as a complementary tool inpremarket device evaluation and postmarket foren-sic investigations. As part of an FDA initiative toevaluate the utility of CFD for assessing medicaldevice safety and to understand the limitations ofCFD in this field, in vitro blood damage testing wasperformed on a simple nozzle model to generateinterlaboratory data for comparison with computa-tional predictions. The focus of this particular studyis not on CFD analytical techniques, but rather onthe importance of the experimental data needed tovalidate computational models. Current limitationsin experimental hemolysis studies have included thefollowing: (i) not accounting for important differ-ences in RBC fragility between species in variousflow regimes (19,20); (ii) not implementing realistictimes of RBC exposure to high stresses (usually lessthan 1 s) during the in vitro testing; (iii) notaddressing how blood collection and test conditionssuch as the source of animal blood, drawing tech-nique, anticoagulation, time after withdrawal, tem-perature, and pH may affect flow-induced hemolysis(8); (iv) not disclosing important details about themodels (e.g., surface roughness and corner con-tours) (21); and (v), failing to adequately verifyexperimental hemolysis results among multiplelaboratories (22).

In this study, three independent laboratories con-ducted in vitro hemolysis experiments on a reversiblenozzle model to relate physical parameters (e.g., flowrate, device geometry, and animal blood properties)to RBC trauma. To our knowledge, this is the firstmultilaboratory study to comparatively evaluate invitro hemolysis in the same specific test model. Amultilaboratory assessment is beneficial in that itdraws on expertise from multiple sources, incorpo-rates real world testing biases, and can provide largesample sizes that may be impractical to obtain froma single center. Complementary interlaboratorystudies that describe the flow field characteristics inthis nozzle model using particle image velocimetrymeasurements (23) and computational simulations(24,25) under slightly different flow conditions arepublicly available at https://fdacfd.nci.nih.gov toencourage further improvements in the developmentand evaluation of medical device blood damagemodels using computational simulation tools.

MATERIALS AND METHODS

Nozzle modelThe FDA benchmark nozzle model, which forces

blood from a tube with a 12-mm inner diameter intoa throat region with a 4-mm inner diameter (40 mmlong), was based on earlier nozzle model designs thatwere shown to induce measurable hemolysis (26,27).The nozzle model contains regions of sudden andgradual changes in cross-sectional area, similar tomany actual blood-contacting medical devices (e.g.,catheters, syringes, hemodialysis tubing, oxygen-ators, and hypodermic needles). Figure 1 displays the

FIG. 1. Nozzle model design (left) andschematic of the recirculating blooddamage test loop (right). The model wasevaluated with a sudden contraction atthe entrance to the throat at flow rates of5 and 6 L/min and with a gradual coneinlet configuration at 6 L/min.

THOUGHTS AND PROGRESS238

Artif Organs, Vol. 39, No. 3, 2015

https://fdacfd.nci.nih.gov

critical dimensions, along with the two flow orienta-tions of the test model (on the left): (i) with a sharp-cornered sudden contraction at the inlet to the nozzlethroat at blood flow rates of 5 and 6 L/min, and (ii)with the model reversed such that the gradual conicalsection was at the inlet to funnel blood into thestenotic throat at 6 L/min. Three identical nozzlemodels were fabricated in blocks of acrylic(2.5 × 2.5 × 16.5 cm) through a process of milling,hand-polishing, vapor-polishing, and temperatureannealing. Since rough surfaces (Ra > 0.6 μm) aremore likely to contribute to hemolysis (13,21,28),smooth blood-contacting surfaces were maintainedthroughout the nozzle model and flow loop. Thesesurfaces were assessed using a stereomicroscope andmetrology rubber (Reprorubber, Flexbar MachineCorp., Islandia, NY, USA), a profile optical com-parator (PH-350, Mitutoyo, Aurora, IL, USA), andan optical surface profilometer (ContourGT-K1,Bruker AXS, Madison, WI, USA). The models con-formed to the following geometrical parameters: (i)an inlet diameter of 12.00 ± 0.13 mm with a throatdiameter of 4.00 ± 0.08 mm; (ii) an angle of suddencontraction of 90.0 ± 0.1°; (iii) a radius of curvature atsudden contraction corner <25 μm; (iv) angle ofgradual cone of 20 ± 1°; (v) misalignment of the conegeometry from the center of the tube <0.03 mm; and(vi) surface roughness Ra < 0.3 μm with no visibletool marks on the blood-contacting surfaces. Pol-ished stainless steel tubes (12-mm inner diameter,15-cm length, Ra < 0.3 μm) were sealed usingpolytetrafluoroethylene thread tape into counter-sinks on both sides of the acrylic model to maintainundisturbed entry and exit flow (∼19 cm long) to andfrom the nozzle throat section. To account for anypossible geometric or surface discrepancies amongthe three nozzle models, each model was tested byeach laboratory.

Test circuitThe flow loop shown in Fig. 1 was standardized to

conduct sensitive and reproducible in vitro hemolysistesting using a magnetically levitated centrifugalblood pump (Levitronix, Waltham, MA, USA) torecirculate the blood. Blood from the pump outletentered a straight section of half-inch-inner-diameterTygon S-50-HL medical grade tubing (St. Gobain,Akron, OH, USA), which connected to the verticallymounted nozzle model. A vertical configuration wasused to assist in circuit priming and to match theprevious velocity characterization experiments(23). Pressure drop was measured across the modelwith digital pressure transducers (Digimano 1000,Netech, Farmingdale, NY, USA, or Model

1502B01EZ5V20GPSI, PCB Piezotronics, Depew,NY, USA). Blood samples were acquired from a low-pressure stopcock port distal to the nozzle outlet. Thereservoir was designed to increase test sensitivity byminimizing blood volume, to promote blood mixingand minimize stagnation zones, and to withstand col-lapse from low pump inlet pressures while maintain-ing a closed system. The reservoir was composed of a500-mL medical-grade PVC bag (Qosina, Edgewood,NY, USA) with a sealable air vent. The bag was cutand clamped over an acrylic reservoir base withblood inlet and exit ports with half-inch outer diam-eters, and it was heat-sealed diagonally to reduce theloop volume and facilitate blood mixing. Eachexperiment used an average blood volume of 275 or295 mL, depending on the test site.

A thin-walled stainless steel tube (10-mm innerdiameter, 50 cm long) enclosed within a waterjacket was used as an intraluminal heat exchangerto maintain a blood temperature of 25.0 ± 1.5°C. Aclamp-on 9XL ultrasonic flow probe and meter(T110R, Transonic Systems, Inc., Ithaca, NY, USA),factory-calibrated with blood and regularly cali-brated on site with phosphate-buffered saline (PBS)or blood, were used to monitor blood flow rate.Lastly, an access port was located at the bottom ofthe loop to allow draining and refilling of the systembetween tests.

Blood preparation and quality assessmentFor hemolysis testing, the three laboratories used

bovine blood obtained from commercial donoranimals or from an abattoir. To determine whetherblood species could affect the results, two of thegroups later repeated a subset of the test conditionsusing porcine blood. Donor blood was drawn fromlive animals via venipuncture using a 12-G lancet andarrived at the testing facility within 24 h of beingdrawn. Abattoir blood was obtained immediatelyafter death using venipuncture as described above orgravity-collected into a bucket after vascular incisionand arrived at the test site within 1.5 h of the draw.Each blood pool was used for two consecutive daysof testing, and all testing was completed within 80 hof the blood being drawn. The blood was stored at4°C when not in use. Each laboratory used a total ofsix distinct blood pools to complete the testing withbovine blood.

Anticoagulation was achieved using a 1:8 ratio ofacid citrate dextrose solution A (ACDA) to blood.ACDA was used in this study due to its ability toeffectively preserve blood cells over multiple days ofstorage (29). For each multiple-day test, 4 L of bloodwas filtered through a latex-free mesh (40–75 μm pore

THOUGHTS AND PROGRESS 239


size) and pooled together at the test facility. Hemato-crit was determined by centrifugation of standardcapillary tubes and adjusted to 36.0 ± 1.5% by eitherhemodilution with PBS or hemoconcentrationthrough plasma removal. The blood was gently mixedprior to loading into the test loop.

Blood quality was assessed by monitoring the base-line hemolysis level of the normalized blood pool andby a rocker bead test, which was used to characterizeRBC mechanical fragility for each separate bloodpool on each day of testing (30,31). For the RBCmechanical fragility test, five 7-mL test tubes(silicone-coated glass; Becton Dickinson Vacutainer,Fisher Scientific, Franklin Lakes, NJ, USA) were uti-lized such that three test tubes contained five stain-less steel ball bearings each (1/8″ diameter, BNMX-2Type 316 SS balls, Small Parts, Miami Lakes, FL,USA), while the remaining two test tubes containedno ball bearings and served as controls. Each tubewas filled with 3 mL of blood, placed on a platformrocker (M79700 Platform Vari-Mix Rocker,Barnstead Thermolyne Corp., Dubuque, IA, USA),and rocked for 1 h at 18 cycles per minute with a 17°oscillatory amplitude. After 1 h, plasma free hemo-globin concentration (fHb) was measured in eachsample. The RBC mechanical fragility index (MFI),shown in Eq. 1, was used as a measure of RBCmechanical fragility (30):

MFIfHb fHbtHb fHb

final control

pool control

= −−

∗100 (1)

where fHbfinal is the plasma free hemoglobin concen-tration (mg/dL) in the samples with beads, fHbcontrol isthe hemolysis level (mg/dL) in the rocked controlsamples without beads, and tHbpool is the total hemo-globin concentration (mg/dL) of the pooled blood.

Blood viscosity was measured each test day at ashear rate of 200 s−1 using a cone-and-plate rheometer(Brookfield, Middleboro, MA, USA) or an oscillatoryviscometer (Vilastic-3, Vilastic Scientific, Austin, TX,USA). Density was calculated by weighing 100 mL ofblood, and plasma protein concentration was mea-sured with a refractometer (TS 400, Reichert, Depew,NY, USA). All three laboratories measured the pHlevels of the blood. Additionally, two laboratoriesmeasured carbon dioxide and oxygen levels with ablood gas analyzer and blood glucose concentrationsusing a glucometer (Hemocue, Glucose 201 Analyzer,Cypress, CA, USA).

Flow conditionsThe Reynolds number (Re) in the throat of the

model was calculated as

Re = 4ρπμ

QD

(2)

where ρ is the density of blood, Q is the measuredblood flow rate, μ is the dynamic viscosity of blood,and D is the 4-mm throat diameter of the model.Initially, our intent was to study blood flow at Reyn-olds numbers ranging from 500 to 6500 to correlatehemolysis levels with (i) the shear stresses estimatedusing particle image velocimetry in similarly preparedacrylic nozzle models (23) and (ii) the hemolysis pre-dictions obtained through computational simulations(24). In the companion particle velocimetry study(23), the authors evaluated flow through the FDAnozzle model for throat Reynolds numbers that cor-responded to a blood flow rate range of 0.4–5.0 L/min.This flow range encompassed both laminar and turbu-lent flow regimes. However, preliminary hemolysistesting with the nozzle model revealed that blood celldamage was minimal when the flow rate was below5 L/min, so low flow rates were excluded fromhemolysis testing. When the flow rate exceeded 6.5 L/min, flow-induced cavitation could be acousticallydetected at the sudden-contraction entrance to thethroat region using a hydrophone (Model 132A32,PCB Piezotronics) with a signal conditioner (Model480C02, PCB Piezotronics). Since differentiationbetween shear-induced blood damage and cavitation-related hemolysis would not be possible, the nozzlemodel was operated below the cavitation thresholdwith testing only performed at 5 and 6 L/min.

Experimental procedureHemolysis testing involved the alternation of the

nozzle model between two identical flow loops thatwere operated simultaneously. For each nozzlemodel experiment, a corresponding control test pre-ceded or followed that matched pressure and flowconditions using the same blood pool, test loop, andblood pump. In a few cases, a single control loopcorresponded to two distinct nozzle-containing testloops. For the control experiments, the acrylic modeland stainless steel extensions were removed andreplaced with two polycarbonate reducers (1/2″ to1/4″), a 30″ looped segment of Tygon S-50-HLmedical grade tubing 1/4″ in inner diameter, and anadjustable hosecock clamp. The baseline control testswere used to compensate for the blood damagecaused by the flow loop components (e.g., the pump,connectors, reservoir, and tubing) other thanthe nozzle. To isolate the blood damage caused bythe nozzle model itself at a specific flow condition,the free hemoglobin values generated by the baseline



control loop were subtracted from the results of thecorresponding nozzle test loop. To differentiatebetween the 6-L/min control loops, the baselinecontrol corresponding to the nozzle conical inletdirection is referred to as “Control A,” and that forthe sudden-contraction entrance is “Control B.”

PBS was recirculated through each of the flowloops for 15 min prior to the beginning of the bloodtests to wet the blood-contacting surfaces. The PBSwas then drained, and blood was gravity-filled fromthe lowest point in the loop to minimize mixing withair. The blood was allowed to circulate for 5 min at aflow rate <2 L/min to de-air the system before thestart of each test. Each experiment lasted 2 h withblood samples drawn every 40 min. One milliliter ofblood was withdrawn from the sampling port anddiscarded before two 2-mL blood samples weredrawn for analysis.

A standardized cleaning protocol was establishedto reduce procedural discrepancies among the labo-ratories. At the start of each test day, new reservoirbags and tubing sets were used. In between same-dayexperiments, the loop was washed twice with freshPBS to remove any visible trace of blood. At the endof each test day, the acrylic nozzle model, stainlesssteel extensions, heat exchanger, pump head, andacrylic reservoir base were rinsed with PBS, washedwith soap/degreaser (Versa-Clean, 10× dilution,Fisherbrand, Pittsburgh, PA, USA; or 1% SimpleGreen, Sunshine Makers, Inc., Huntington Beach,CA, USA) and water, then washed with 70% isopro-panol or ethanol, rinsed with deionized water, andallowed to completely dry before reassembly withnew tubing. No metal brushes were used during thecleaning process to avoid scratching the blood-contacting surfaces. Pump heads were replaced after2 consecutive days of testing.

Hemolysis assays and analysisTo prepare plasma for analysis, blood samples

were first centrifuged at no more than 2000 × g for15 min, followed by plasma isolation andrecentrifugation at 13 000 × g for 15 min. Using theresulting supernatant plasma, hemolysis was deter-mined spectrophotometrically by measuring thechange in fHb. The Cripps method (32), used by allthree laboratories to quantify hemolysis, is based onthe oxyhemoglobin levels at three wavelengths (560,576.5, and 593 nm) as shown below (33):

fHb mg dL conv

dilnmnm nm

{ } = ( )

∗ −+⎡

⎣⎢⎤⎦⎥

∗( )AA A

576 5560 593

2.

(3)

where ‘conv’ refers to the spectrophotometerconversion factor and ‘dil’ is the dilution factor of thesample. The conversion factor, determined bycalibration of the spectrophotometer with serialdilutions of hemoglobin standards, was approxi-mately 177.6 for all of the spectrophoto-meters used in this study (33). To ensure thatmeasurements were made within the linear range ofthe spectrophotometer, fHb values above 150 mg/dLwere diluted with PBS and re-measured. Prior toconducting the nozzle hemolysis study, stock fHbsolutions were distributed, and all laboratoriesmeasured within ±3.5 mg/dL of target values rangingfrom 0–100 mg/dL (i.e., <3.6% coefficient ofvariation). As multiple blood samples were assayedover the 2-h loop tests, all ΔfHb values were based ona linear regression analysis of the slope of the fHbversus time plots using eight measurements takenover four sampling times.

To account for variations in the test parameters(e.g., flow rate, blood volume, hematocrit, and totalblood hemoglobin concentration) between experi-ments, the measured fHb was normalized by calcu-lating the normalized index of hemolysis (NIH) (22).

NIH g L fHbHct

100100

1001{ } = −Δ * * **

VQ t

(4)

where V is the volume of blood in the flow system(L), Q is the blood flow rate (L/min), Hct is thehematocrit of the blood (%), and t is the duration ofthe test (min). The modified index of hemolysis(MIH) can be derived from the NIH and accounts forthe total hemoglobin concentration in the blood(tHb, mg/dL) (8).

MIH mg mg NIHtHb

{ } = × 106

(5)

MIH is reported in dimensional units of milligrams ofhemoglobin released into plasma divided by the mil-ligrams of total hemoglobin pumped through the loop(6). To minimize decimal places, a constant factor of106 is used (8). To account for the hemolysis causedsolely by the nozzle model (MIHnozzle), the MIH valuemeasured in the matched control loop (MIHcontrol) wassubtracted from the blood damage caused by thenozzle test loop (MIHtest) according to Eq. 6.

MIH MIH MIHnozzle test control= − (6)

Sample size and statistical analysisHemolysis data were collected on two adjacent

days for each blood pool, resulting in an uneven



number of repeats for the three test conditions. Spe-cifically, the sudden-contraction nozzle inlet condi-tion at 6 L/min had 36 total repeats, whereas only 27repeats were collected for each of the other twonozzle test conditions. The number of repeats wassufficient to assess experimental reproducibility(including intraday repeatability) and the effect ofblood aging between test days.

Statistical analyses were performed using unpairedt-tests, and a P value of less than 0.05 was consideredstatistically significant. When multiple conditionswere compared, an F-test was first performed todetect differences between groups. Linear regressioncoefficients were used to quantify the variability ofthe results and the linear association of the hemolysisdata with the test duration and other test parameters.The standard deviation (SD) and percentage coeffi-cient of variation (% CV = SD/mean times 100%)were reported to indicate the variability of the resultsat a given condition. Grubbs’ extreme studentizeddeviate test was used to detect outliers in the dataset. Additionally, to estimate the contribution ofviscosity, flow rate, and density measurement uncer-tainties towards the overall variability in thehemolysis data, an error propagation analysis wasperformed using commercial CFX software(ANSYS, Inc., Canonsburg, PA, USA) and a modi-fied Grigioni (Lagrangian) blood damage model (34)with Giersiepen–Wurzinger constants (35) inMATLAB (MathWorks, Natick, MA, USA).

RESULTS

The chemical and physical properties of bovineand porcine blood after hematocrit adjustment to36% are shown in Table 1. In general, the measuredblood properties were similar for each laboratory andindependent of the blood source (i.e., live donor orabattoir), method of blood collection (i.e., venipunc-ture vs. bucket), and storage age of the blood at thestart of each experiment (i.e., 3–80 h old). While thedaily order in which the experiments were performed

was inconsequential, the blood storage time (e.g.,first day of testing vs. second day) did have a signifi-cant effect on the dynamic hemolysis test results.When the same test condition was run on consecutivedays, hemolysis levels were higher on the second testday compared with the first by about 20% (n = 18,P < 0.03). In spite of this difference, the hemolysisresults for the three laboratories have been collatedbased on test condition irrespective of test day, aseach test condition was evaluated an equal number oftimes on both days.

The effects of device geometry and flow rate onhemolysis were studied under three test conditions:(i) sudden-contraction nozzle inlet at 5 L/min, (ii)gradual cone inlet at 6 L/min, and (iii) sudden-contraction inlet at 6 L/min. The hemodynamic oper-ating conditions for each lab were comparable withmean blood flow rates, all being within 0.2 L/min, or∼4%, of the target values of 5 and 6 L/min. The mea-sured pressure drops across the nozzle models werealso similar among the laboratories (5–7% CV).Pressure drop variations among the laboratorieswere attributed to pump speed discretization(±100 rpm) and to calibration differences in the flowand pressure probes. Table 2 shows that the targetflow rates of 5 and 6 L/min equated to turbulentnozzle throat Reynolds numbers of 6650 ± 570 and7950 ± 660, respectively. At both flow rates, the flowin the half-inch-inner-diameter tubing of the loopwas transitional in nature (Re = 2220 ± 190 and2670 ± 220 for 5 and 6 L/min, respectively). Based onmean velocities, the average exposure time of anRBC to the throat region was 5 to 6 ms for the 6- and5-L/min flow rates, respectively.

The average initial fHb value at the start of alldynamic hemolysis tests with bovine blood was15.5 ± 7.3 mg/dL. During hemolysis testing, the fHbincreased linearly over the duration of the experi-ment as shown in Fig. 2 (e.g., r2 > 0.94 for sudden-contraction nozzle inlet at 6 L/min for the 12repeats from one laboratory). The fHb magnitudeand linearity indicate that a 2-h experimental run

TABLE 1. Physical and chemical properties of the bovine and porcine blood adjusted to 36% hematocrit

Bloodspecies

Originalhematocrit

(%)

Adjustedhematocrit

(%)Density(g/mL)

Viscosity at25°C (cP)‡

Plasmaprotein(g/dL)

Totalhemoglobin

(g/dL)Glucose(mg/dL)

pH at37°C

pO2

(mm Hg)pCO2

(mm Hg)Blood

fragility

Bovine* 38.0 ± 2.2 36.2 ± 1.1 1.04 ± 0.01 4.21 ± 0.32 6.5 ± 0.6 11.9 ± 0.8 365 ± 59 6.82 ± 0.10 71 ± 23 87 ± 21 0.54 ± 0.15Porcine† 44.4 ± 2.9 36.4 ± 0.8 1.04 ± 0.01 4.06 ± 0.58 6.8 ± 0.6 11.6 ± 0.9 — 6.81 ± 0.10 56 ± 32 86 ± 10 0.42 ± 0.06

Values are shown as mean ± SD. * Values for bovine blood were based on data taken from three laboratories (n = 36 for all parametersexcept plasma protein, glucose, and blood gases, for which n = 18 to 28). † Porcine blood properties were measured in two laboratories(n = 16 for all parameters except glucose and blood gases, for which n = 4 to 12). The glucose concentration in porcine blood anticoagulatedwith acid citrate dextrose solution A was not reported, as it often exceeded the measurable range of the glucose analyzer. ‡ Viscosity ofbovine and porcine blood was measured at a shear rate of 200 s−1.



time was sufficient to generate blood damage withthe nozzle model and that autohemolysis was not alikely factor during the tests. However, Fig. 2 alsoillustrates the variability in hemolysis test resultswhen 12 experiments were performed on differentdays with different pools of blood; the variability inslope of the hemolysis versus time plots was 47%CV.

Multilaboratory changes in fHb concentration forthe three test conditions and their respective controlsare displayed in Fig. 3. In all cases, the hemolysisvalues were above the minimal detection level of1–2 mg/dL for the fHb concentration assay (33). Forthe 5- and 6-L/min sudden-contraction nozzle inletflow conditions, the hemolysis levels were two to ninetimes greater than in their corresponding controlloops. In contrast, the gradual cone nozzle inlet testcondition at 6 L/min did not generate significantlymore hemolysis than the control loop. This resultindicates that the gradual cone inlet and the 4-mmthroat of the nozzle produced minimal damage to

RBCs at the highest tested flow rate. The two barsdenoted by symbols in Fig. 3 each contain a singledata outlier as specified by Grubbs’ test. These out-liers were likely caused by procedural errors or bloodcontamination and were not included in the subse-quent analysis and histograms (Fig. 4).

The flow-induced hemolysis caused only by thenozzle (MIHnozzle) was calculated by subtracting theblood damage generated in the correspondingcontrol loop. Figure 4A shows the MIHnozzle resultsfor each test condition and laboratory. There were nostatistical differences among the hemolysis levelsmeasured at the three laboratories when the sudden-contraction geometry was at the nozzle inlet at 5 and6 L/min (P > 0.09). However, for the test conditionthat caused the least amount of hemolysis (gradualcone inlet condition at 6 L/min), Laboratory 3 gener-ated a higher level of hemolysis than Laboratory 2(P = 0.03). The low MIHnozzle values for the gradualcone inlet indicate that this nozzle configuration gen-erates comparable hemolysis to the correspondingcontrol loop.

TABLE 2. Flow parameters in the nozzle model for the three test conditions

Sudden-contraction inlet,5 L/min (n = 26)

Gradual cone inlet,6 L/min (n = 26)

Sudden-contraction inlet,6 L/min (n = 36)

Measured flow rate (L/min) 5.05 ± 0.12 5.94 ± 0.14 6.07 ± 0.08Pressure drop across nozzle model (mm Hg) 217 ± 15 309 ± 22 297 ± 15Pump speed (rpm) 3420 ± 70 4020 ± 110 3970 ± 60Reynolds number in nozzle throat 6650 ± 570 7860 ± 640 8020 ± 670Reynolds number in half-inch-inner-diameter tubing 2220 ± 190 2630 ± 210 2670 ± 220

Values are shown as mean ± SD for three laboratories combined.

FIG. 2. Changes in plasma free hemoglobin concentration forbovine blood during 120 min of testing for Laboratory 1, showingall 12 individual tests of the 6-L/min sudden-contraction nozzleinlet condition (r2 > 0.94 for each individual test). The bold linerepresents the mean ± SD values at each blood sample timepoint, and similar trend lines indicate which experiments used thesame blood pool on 2 consecutive test days.

FIG. 3. Change in plasma free hemoglobin concentration forbovine blood after 120 min of testing in nozzle and control con-ditions for three laboratories. Values are shown as mean ± SD foreach laboratory, and the number of replicates per lab (n) isindicated for each test condition. §A single outlier was present andincluded in the specified data set.



The collated hemolysis values from the threelaboratories for each nozzle test condition weresignificantly different from one another (Fig. 4b,P < 0.001). Despite having similar nozzle throatReynolds numbers and afterload pressures at a flowrate of 6 L/min (Table 2), the sharp corner geometryof the sudden-contraction inlet produced much morehemolysis (MIHnozzle = 1.239 ± 0.667, n = 36) than thegradual cone inlet condition (MIHnozzle = 0.021 ±0.128, n = 26). Flow rate was also a major factor inhemolysis generation; the sudden-contraction nozzleinlet caused more hemolysis at 6 L/min than at 5 L/min (MIHnozzle = 0.292 ± 0.249, n = 26). The ratioof hemolysis values between the test conditions(mean ± SD) was also calculated. Relative to the 5-L/min sudden-contraction inlet condition, the 6 L/minsudden-contraction inlet generated on average4.24 ± 0.79 times more hemolysis, while the 6-L/minconical inlet generated approximately 14 times less(i.e., ratio = 0.07 ± 0.09).

The same relative trends in hemolysis generationfor each test condition were observed independent ofparameter or index (ΔfHb, NIH, MIH). There weresignificant Pearson product-moment correlationcoefficients (r) for ΔfHb versus NIH (0.994), ΔfHbversus MIH (0.988), and NIH versus MIH (0.997).

Three types of variability associated with the invitro hemolysis testing were determined in thisstudy: (i) repeatability—a comparison of identicaltests performed on the same day with the same testloop and using the same blood pool; (ii)intralaboratory variability—variability at each testfacility when performing the same test with differ-ent blood pools on different days; and (iii)interlaboratory variability—a comparison of the dif-ferences in hemolysis values across the three testfacilities. The repeatability for the sudden-contraction nozzle inlet hemolysis tests at both 5and 6 L/min, in terms of the percent coefficient ofvariation, was 18% (n = 14). Some of the resultsused to assess repeatability were collected duringpreliminary reproducibility testing of the models.The intralaboratory variability of the MIHnozzle

values for the three laboratories ranged from 47 to94% (from Fig. 4A). Hence, the inherent variabilityassociated with repeat experiments on a single daywas significantly less than the intralaboratory vari-ability established over multiple days of testing withdifferent blood pools. Additionally, it was deter-mined that intralaboratory variability was greaterthan the interlaboratory variability (54–85%) usinga random-effect model (F > 100, P < 0.001).

To estimate the contribution of flow-relateduncertainties to the overall variability in thehemolysis results, an error propagation analysis wasperformed post hoc. We estimated that variability inviscosity, flow rate, and density measurements couldlead to a 23% uncertainty in the Reynolds number.After determining the viscous and Reynolds stressesin the nozzle using a CFD simulation, a modifiedGrigioni (Lagrangian) blood damage model (34)was used to predict that the variability in these flow-related parameters could only account for up to15% CV in the hemolysis index at each test condi-tion. As the intra-laboratory hemolysis variabilityranged from 47 to 94% CV, this indicates that thecombined uncertainty associated with the experi-mentally measured flow rate, viscosity, and, to alesser extent, density accounted for less than a thirdof the overall intra- and interlaboratory variabilityin hemolysis.

The RBC rocker bead mechanical fragility testwas used to determine whether the MFI of eachblood pool correlated with the hemolysis produced

FIG. 4. MIH values for the three nozzle test conditions aftersubtraction of the control loop MIH values (A) for each laboratoryand (B) averaged for the three laboratories. ‡A single outlierwas removed from the specified data set (n = total number ofreplicates).



by the nozzle. For each of the three laboratories, themean RBC MFI values for 36% hematocrit bovineblood were 0.56 ± 0.10, 0.45 ± 0.12, and 0.62 ± 0.18,which were significantly different from one another(P < 0.02). With a coefficient of variation of only 4%(n = 30), the repeatability of the RBC mechanicalfragility tests using the same blood on the same testday at each laboratory was very high. Theintralaboratory variability (% CV) ranged from 17 to29% and the interlaboratory variability was 28% forthis test, and there was no significant difference inMFI between the first and second test days(P > 0.05). Overall, there was not a clear correlationbetween the MFI of the blood on each test day andthe amount of hemolysis generated by the three sepa-rate nozzle test conditions (P > 0.05).

A subset of experiments was also conducted bytwo laboratories using both porcine and bovineblood to determine if blood species affectedhemolysis generation in the MFI test and the nozzlemodel. As there was no interlaboratory differencein the nozzle hemolysis results (P > 0.08), the datawere collated for the two laboratories. PorcineRBCs better match the rheological and morphologi-cal properties of human RBCs, including a largermean corpuscular volume, in comparison to bovineblood cells (12,36). The RBC mechanical fragilitytests showed that bovine blood had a higher meanMFI than porcine blood (0.54 ± 0.15 vs. 0.42 ± 0.06,respectively; P < 0.01; Table 1), perhaps due to aslightly higher concentration of protective totalprotein in the porcine blood (37). While the MFItest indicated that bovine blood may be more sen-sitive to damage, Fig. 5 shows that there was no sta-tistical difference in flow-induced hemolysisbetween porcine and bovine blood (both at 36%

Hct) using the FDA nozzle model with the sudden-contraction inlet at flow rates of 5 and 6 L/min(P = 0.57). There was, however, a statistical differ-ence between the species for the relative ratio ofhemolysis generated at the 6-L/min versus the 5-L/min condition (P = 0.03). The hemolysis ratio was8.93 ± 2.07 for porcine blood and 4.02 ± 1.00 forbovine blood.

DISCUSSION

As in vitro hemolysis test results are a function ofmany different factors such as blood species, bloodmanagement, temperature, and test model (8,38,39)and can be laboratory-dependent, this multi-laboratory study was conducted to evaluate in vitrohemolysis testing techniques and to generate experi-mental data to assist in the implementation of CFD forthe blood damage safety evaluation of medicaldevices. In previous studies, similar nozzle models andtest conditions were used (i.e., bovine blood tested at5 or 6 L/min through 10–11-mm-diameter inlet tubespast sharp-cornered and tapered inlets into constrict-ing 4–5-mm-diameter throats that were 12–15 mm inlength), but the hemolysis results differed by a factorof approximately 50 (26,27). This discrepancy reflectsthe difficulty in comparing in vitro hemolysis testresults across different studies when there is a paucityof repeated tests and differences in temperature,pump afterload, pump operation, blood volume, flowrate, loop components, hematocrit, blood species,blood drawing methods, and anticoagulation. Asanother example, literature values of NIH for theBiomedicus BP-80 pump (Medtronic, Minneapolis,MN, USA) tested under similar operating conditionsrange from 0.0007 to 0.018 g/100 L, with individualstudy coefficients of variation ranging from 18 to 94%(40–42). Advantages of the current study are the fol-lowing: (i) the nozzle model was comparatively testedat three different laboratories and had similar pres-sure losses and hemolysis results, (ii) many test repli-cates were performed using both bovine and porcineblood, (iii) the flow-induced nozzle hemolysis levelwas obtained by correcting for the backgroundhemolysis caused by the pump and loop components,and (iv) all testing and nozzle model parameters areprovided in this text.

As expected, the sudden-contraction inlet configu-ration caused significantly higher hemolysis than thegradually tapering cone inlet, suggesting that thesharp corner geometry of the sudden-contractioninlet was the primary source of blood damage. Ashemolysis testing was performed below the thresholdfor cavitation formation in this study, any measured

FIG. 5. Comparison of the MIH between bovine and porcineblood for two nozzle test conditions after subtraction of the controlloop MIH values at two laboratories. Values are shown asmean ± SD (n = total number of replicates per test condition).



RBC lysis was attributed to flow-generated stresses.As shown by CFD analysis, the sudden-contractionconfiguration produced high viscous shear stresses atthe sharp corner and large variations in the wall shearstress magnitudes and distributions (25). Further-more, the wall shear stresses associated with thegradual cone inlet condition were smaller than at thesudden-expansion outlet and did not cause muchhemolysis (24,25). The parameters used in this studyto report hemolysis (i.e., plasma free hemoglobin,NIH, and MIH) did not impact data interpretationbecause the blood flow rate, hematocrit and totalhemoglobin levels, blood volume, and sampling timewere carefully controlled. Therefore, irrespective ofwhich hemolysis parameter is used, the results dis-played in Fig. 4B (e.g., MIH) may be used to helpdevelop and verify predictive CFD models of blooddamage as long as the variability in the hemolysisresults is also taken into consideration. However,considering that blood damage test results may varydue to test temperature, length between blood col-lection and use, anticoagulation, blood pH, oxygen-ation, and glucose levels (8), it should be noted thatthe hemolysis results reported here are specific forthe blood parameters used in our testing (i.e.,passive-flow nozzle model with a 4 mm throat, bovineand porcine blood cold-stored in some cases for up to80 h after withdrawal, ACDA anticoagulation, bloodtest temperature of 25°C, 36% hematocrit).

Another important aspect of this study was toexamine the variability associated with in vitrohemolysis experiments. To account for the high vari-ability in the test results and to conduct robust statis-tical analyses, a large number of replicates werenecessary. By utilizing three laboratories, a com-bined total of 26–36 data points were accumulated ateach nozzle test condition. While identical clinicalblood pumps were used and a standard operatingprocedure was followed, subtle differences in experi-mental technique (including blood collection, han-dling and sampling), blood source and age, animal-to-animal variation, equipment calibration, variableloop and reservoir configurations, and cleaningprocedures still may have contributed to theinterlaboratory variability shown in Fig. 3. Addition-ally, much of the intralaboratory variability could beattributed to inherent differences in blood cell fragil-ity not captured by the rocker bead fragility test,performance changes of the individual flow loopcomponents, de-airing inconsistencies, loop con-struction alterations, or variations in operating thesystems. In this study, some day-to-day variabilitywas compensated for by subtracting the hemolysismeasured in a paired control flow loop (Fig. 4). This

was important as the parallel tests showed a signifi-cant positive correlation between the hemolysiscaused by the control loop and its correspondingnozzle test loop (P < 0.002).

A comparison between bovine and porcine bloodwas performed using the RBC mechanical fragility(rocker bead) test and hemolysis tests of two nozzleflow conditions (by two laboratories), as both bloodtypes are readily available in large quantities andoften used for in vitro medical device testing. Animportant area of investigation is determining how invitro hemolysis results obtained with animal bloodcan be extrapolated to human clinical situations.Porcine blood is thought to be more relevant thanbovine blood for in vitro blood damage testing (12).The differences in RBC size and fragility of bovineand porcine blood are likely to be more noticeable insmaller-scale models, such as orifice jet flow modelsor shearing devices with small gaps (12,39), as only asmall percentage of the RBCs may contact the wallsor be exposed to the elevated shear stresses at thesudden-contraction inlet during a single pass (25).

The RBC mechanical fragility tests (with rockedbeads) varied between the laboratories but indicatedthat bovine RBCs were more fragile than porcineRBCs (Table 1). The sudden contraction nozzle inletresults showed no significant difference in RBCdamage between bovine and porcine blood at the 5-and 6-L/min conditions (Fig. 5), yet the relative ratioof hemolysis between flow rates suggests otherwise(P = 0.03). This discrepancy should not be miscon-strued as proof that porcine blood was more sensitivethan bovine blood for the nozzle model. For the 5-L/min sudden-contraction inlet condition, bovine bloodactually yielded a higher average hemolysis value thandid porcine blood. The relative ratio calculations wereunduly impacted by the relatively low blood damagelevels at 5 L/min for both species. Hence, to properlydetermine differences in blood fragility between dif-ferent species, more testing conditions with a widerrange of hemolysis values are required.

The RBC mechanical fragility test also allowed forthe assessment of test variability in a simple systemthat did not include a pump or recirculating flow loop.The RBC mechanical fragility test results were highlyrepeatable (4% CV) when bovine blood from thesame pool was used, with individual intralaboratoryvariability about half that of the nozzle experiments.The statistically significant difference in bovine RBCMFI values among laboratories is likely a combina-tion of both blood- (source, collection method) andlaboratory-specific variability. While a clear correla-tion was not observed between RBC MFI and flow-induced hemolysis, the rocker bead test can still be



useful as a measure of quality control within labora-tories to identify highly fragile blood cells prior totesting (43). It should be noted that the mechanismsfor blood damage in the RBC mechanical fragility test(with laminar flow conditions and multiple bead-surface collisions) should be markedly different thanfor the turbulent flow conditions within the experi-mental nozzle recirculating flow loop.

CONCLUSIONS

A multilaboratory in vitro study was conducted togenerate hemolysis and pressure loss data for abenchmark nozzle model. The hemolysis levels forthe three test conditions were similar across the inde-pendent laboratories, suggesting that the majorfactors affecting red blood cell damage were con-trolled in the study. Despite high intralaboratoryvariability, statistically significant differences amongthe three test conditions were observed, with thesudden-contraction nozzle entrance causing the mostblood damage. Because the hemolysis experimentswere performed using a well-defined model, theseresults may help to support, validate, and advancethe development of predictive computational modelsof blood damage, as long as the dependences on thespecific blood parameters and variability of thehemolysis results are taken into consideration.Further studies are needed on other models anddevices over a range of operating conditions toprovide the much-needed hemolysis data that can beused to augment computational predictions of blooddamage.

Acknowledgments: This study was supported bythe US Food and Drug Administration’s CriticalPath Initiative. From the FDA, we would like tothank Dr. Meijuan Li for providing invaluable statis-tical analyses, Dr. Prasanna Hariharan and GavinD’Souza for performing computational simulationsand error propagation analyses, Jean Rinaldi forreviewing the manuscript, and Drs. Sandy Stewartand Qijin Lu for technical support. We recognize andappreciate the support of Brittany Arcuri, ChristineFlick, and Elizabeth Endrizzi for assisting withexperiments at the Cleveland Clinic, and AndrewWearden and Charles Lutzow for helping withexperimental data collection at the University ofPittsburgh. We would also like to acknowledge theefforts of Dr. Steven Day, Matthew Giarra, and AlexShip at the Rochester Institute of Technology in fab-ricating the nozzle models. Lastly, the authors wouldalso like to thank Dr. Kurt Dasse, Farzad Parsaie,and the team at Levitronix LLC for providing cen-

trifugal blood pumps for this study through aMaterial Transfer Agreement. Any mention ofcommercial products and/or manufacturers does notimply endorsement by the US Department of Healthand Human Services.

Conflict of Interest: The authors declare that theyhave no conflicts of interest.

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