ISO 17853:2010
(Main)Wear of implant materials — Polymer and metal wear particles — Isolation and characterization
Wear of implant materials — Polymer and metal wear particles — Isolation and characterization
ISO 17853:2010 specifies methods for sampling wear particles generated by joint implants in humans and in joint simulators. It specifies the apparatus, reagents and test methods to isolate and characterize both polymer and metal wear particles from samples of tissue excised from around the joint implant, obtained at revision surgery or post mortem, and from samples of joint simulator test fluids. Some of these procedures could certainly be adapted for isolation and characterization of particles from human biological fluids (e.g. synovial fluid).
Usure des matériaux d'implant — Particules d'usure des polymères et des métaux — Isolation, caractérisation et quantification
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Standards Content (Sample)
INTERNATIONAL ISO
STANDARD 17853
Second edition
2010-06-15
Wear of implant materials — Polymer and
metal wear particles — Isolation and
characterization
Usure des matériaux d'implant — Particules d'usure des polymères et
des métaux — Isolation, caractérisation et quantification
Reference number
ISO 17853:2010(E)
©
ISO 2010
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ISO 17853:2010(E)
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ISO 17853:2010(E)
Contents Page
Foreword .iv
Introduction.v
1 Scope.1
2 Terms and definitions .1
3 Principle, reagents and apparatus.1
3.1 Principle.1
3.2 Reagents.2
3.3 Apparatus.2
4 Methods of sampling and analysis of polymer and metal wear particles from tissue
samples .3
4.1 Storage and preparation of samples .3
4.2 Procedure for polymer particle isolation .4
4.3 Procedure for metal particle isolation.5
4.4 Collection of particles.6
4.5 Particle size and shape characterization .7
4.6 Particle identification .8
5 Methods of sampling and analysis of polymer and metal particles from joint simulator
lubricants.9
5.1 General .9
5.2 Procedure for polymer materials [e.g. UHMWPE and polyetheretherketone (PEEK)] .9
5.3 Procedure for metal particles.10
5.4 Procedure for ceramic particles .13
6 Test report.13
Bibliography.15
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ISO 17853:2010(E)
Foreword
ISO (the International Organization for Standardization) is a worldwide federation of national standards bodies
(ISO member bodies). The work of preparing International Standards is normally carried out through ISO
technical committees. Each member body interested in a subject for which a technical committee has been
established has the right to be represented on that committee. International organizations, governmental and
non-governmental, in liaison with ISO, also take part in the work. ISO collaborates closely with the
International Electrotechnical Commission (IEC) on all matters of electrotechnical standardization.
International Standards are drafted in accordance with the rules given in the ISO/IEC Directives, Part 2.
The main task of technical committees is to prepare International Standards. Draft International Standards
adopted by the technical committees are circulated to the member bodies for voting. Publication as an
International Standard requires approval by at least 75 % of the member bodies casting a vote.
Attention is drawn to the possibility that some of the elements of this document may be the subject of patent
rights. ISO shall not be held responsible for identifying any or all such patent rights.
ISO 17853 was prepared by Technical Committee ISO/TC 150, Implants for surgery, Subcommittee SC 4,
Bone and joint replacements.
This second edition cancels and replaces the first edition (ISO 17853:2003) and ISO 17853:2003/Cor.1:2004,
which have been technically revised.
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ISO 17853:2010(E)
Introduction
The biological responses to wear particles contribute to the failure of joint implants through bone resorption
and consequent implant loosening. A standardized method of particle retrieval from the tissues followed by
particle characterization is necessary for a uniform approach to wear particle effect investigations. The
characterization of the particles generated from implants in joint simulators also provides valuable information
on the wear properties and performance of the implant being studied.
In the protocols included in this International Standard, for isolation and characterization of particles from both
tissues or test fluids from joint simulators, the particles are isolated and then dispersed using filtration or
embedding in resin for scanning electron microscopy (SEM) or transmission electron microscopy (TEM)
analysis. An alternative protocol for isolation and characterization of metal particles from implants tested in
joint simulators has recently been developed in which the particles are deposited on to wafers for SEM
analysis, without filtration or embedding; see Reference [1]. At the time of writing this International Standard,
this alternative method had not been tested for isolation and characterization of particles from tissues and no
direct comparison between the different methods is currently available. Therefore, the latter method has not
been included in detail.
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INTERNATIONAL STANDARD ISO 17853:2010(E)
Wear of implant materials — Polymer and metal wear
particles — Isolation and characterization
1 Scope
This International Standard specifies methods for sampling wear particles generated by joint implants in
humans and in joint simulators. It specifies the apparatus, reagents and test methods to isolate and
characterize both polymer and metal wear particles from samples of tissue excised from around the joint
implant, obtained at revision surgery or post mortem, and from samples of joint simulator test fluids. Some of
these procedures could certainly be adapted for isolation and characterization of particles from human
biological fluids (e.g. synovial fluid).
The methods given in this International Standard do not quantify the level of wear the implant produces;
neither do they determine the amount of wear from any particular surface. This International Standard does
not cover the biological effects of wear particles or provide a method for evaluation of biological safety.
2 Terms and definitions
For the purposes of this document, the following terms and definitions apply.
2.1
polymer wear particles
particles generated from the wear of polymeric components of an implant
2.2
metal wear particles
particles and particulate corrosion products generated from the wear of metal components of an implant
2.3
ceramic wear particles
particles generated from the wear of ceramic components of an implant
3 Principle, reagents and apparatus
3.1 Principle
Particles of polymeric and metal wear are isolated from tissue samples and simulator lubricants by digestion.
The yield of each particle species is then purified by eliminating any remaining organic debris.
NOTE The methods involved in polymer and metal particle isolation are different and are described in 4.2 and 4.3,
respectively.
The particles are collected, and are characterized and counted (where applicable) using scanning electron
microscopy (SEM) or transmission electron microscopy (TEM).
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ISO 17853:2010(E)
3.2 Reagents
During the analysis, unless otherwise stated, use only reagents of recognised analytical grade and distilled
water or water of equivalent purity.
All reagent solutions shall be filtered through a filter of 0,2 µm or smaller pore size prior to use, to avoid
contamination of the sample by extraneous particles.
3.2.1 Absolute ethanol.
3.2.2 Acetone, 100 % or diluted with distilled water, with a volume fraction of 80 % acetone.
3.2.3 Distilled water.
3.2.4 Fixative, e.g. formalin, diluted with distilled water, with a volume fraction of 10 % formalin.
3.2.5 Hydrochloric acid solution, HCl, c = 0,01 mol/l.
3 3
3.2.6 Isopropanol-water mixtures, ρ = 0,96 g/cm and ρ = 0,90 g/cm .
3.2.7 Papain solution, at 4,8 U/1,5 ml of 250 mM sodium phosphate buffer containing 25 mM
ethylenediaminetetraacetic acid solution (EDTA), pH 7,4.
3.2.8 Sodium phosphate buffer, at 250 mM containing 25 mM of EDTA, pH 7,4.
3.2.9 Proteinase K, 2 g/ml of 50 mM tris(hydroxymethyl)-aminomethane-HCl (TRIS-HCl), pH 7,6.
NOTE For particles isolated from joint simulator serum lubricant, the quantity should be adjusted depending on the
serum percentage of the lubricant and initial serum volume from which particles are isolated. See 5.3.2.
3.2.10 Resin, epoxy, such as EMbed 812.
3.2.11 Sodium dodecyl sulfate (SDS), 2,5 g/100 ml solution in distilled water or 3 g/100 ml solution in 80 %
acetone.
3.2.12 Sodium hydroxide, NaOH, solutions and pellets, c = 0,1 mol/l and 5 mol/l.
3 3 3 3 3
3.2.13 Sucrose solutions, ρ = 1,35 g/cm , 1,17 g/cm , 1,08 g/cm , 1,04 g/cm and 1,02 g/cm .
3.2.14 Tris-hydrochloride buffer, TRIS-HCl, at 50 mM, pH 7,6.
3.3 Apparatus
All apparatus shall be cleaned and triple rinsed with distilled water previously filtered through a filter of 0,2 µm
pore size (3.3.6) before use to remove any contaminating particles.
3.3.1 Aluminium stub.
3.3.2 Balance, with an accuracy of at least 0,1 mg.
3.3.3 Carbon stickers.
3.3.4 Centrifuge tubes, different sizes.
3.3.5 Centrifuge.
3.3.6 Filters, with a pore size of 0,2 µm for filtering reagents and distilled water.
3.3.7 Filtration unit.
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ISO 17853:2010(E)
3.3.8 Formvar-coated copper grids, of 200 mesh size for TEM analysis.
3.3.9 Fourier Transform Infrared (FTIR) Spectroscope.
3.3.10 Heating plate.
3.3.11 Lint-free cloth.
3.3.12 Pipettes, micropipettes and tips.
3.3.13 Polarizing light microscope.
3.3.14 Polycarbonate membrane filters, of pore sizes 10 µm, 1 µm, 0,1 µm, 0,05 µm and 0,015 µm, for
collecting particles.
3.3.15 Scanning electron microscope, SEM, with an energy dispersive X-ray analysis (EDXA) module.
3.3.16 Sterile Petri dishes, with lids.
3.3.17 Syringe, with wide-bore needle.
3.3.18 Teflon-glass potter tissue grinder.
3.3.19 Transmission electron microscope, TEM, with an energy dispersive X-ray analysis (EDXA) module.
3.3.20 Ultrasonic cell disrupter, equipped with a titanium microprobe.
3.3.21 Ultrasonic bath.
3.3.22 Water bath, agitating temperature controlled.
4 Methods of sampling and analysis of polymer and metal wear particles from
tissue samples
4.1 Storage and preparation of samples
Store the tissue at −70 °C (or lower) in a freezer, or at room temperature in a fixative such as formalin (3.2.5),
diluted with distilled water (3.2.4), with a volume fraction of 10 % formalin. Thaw the tissue, if applicable, and
rinse it thoroughly in distilled water before continuing with the extraction method. Remove excess water from
the rinsed tissue by blotting with a lint-free cloth (3.3.12).
Unfixed tissue should be handled under universal conditions.
The nature of surgical instruments used for sample retrieval should be recorded in case of contamination.
NOTE Sampling variability due to specimen origin can occur.
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4.2 Procedure for polymer particle isolation
4.2.1 Tissue digestion
There are many published methods for polyethylene particle isolation from periprosthetic tissues. The
[2] [3] [4]
methods presented here are based on those of Campbell et al. , Tipper et al. and Richards et al. .
Cut the tissue into smaller pieces using a scalpel and blade before digestion to speed up the digestion times.
Extract the lipids from the minced tissue by placing into a 2:1 (volume ratio) chloroform:methanol mixture for
24 h or until the tissue sinks to the bottom of the container. Remove and rinse the tissue with PBS (3.2.8).
Add 5 mM NaOH (3.2.12) to the tissue in a ratio 10 ml of 5 mol/l NaOH to 1 g of tissue and leave to digest for
a minimum of 24 h in an agitating water bath (3.3.22) at 65 °C. Digestion can be judged to be complete when
no visible solid pieces of tissue remain in the suspension.
4.2.2 Purification of the polymer particle yield
4.2.2.1 General
The polymer particles can be purified from the digested tissue in a number of ways. Use one of the methods
described in 4.2.2.2 or 4.2.2.3.
4.2.2.2 Purification of polymer particles by high-speed centrifugation
This method enables all particle sizes to be collected from the nanometre-size range to several millimetres in
length, enabling the total wear volume of particles to be isolated. Cool the digested tissue to 4 °C. Add an
equal volume of ice-cold absolute ethanol (3.2.1). At this point, salts might precipitate. If this is the case, add
ultrapure water until the salt dissolves. Incubate the solution at 4 °C overnight with stirring. Centrifuge the
solution at 20 000 g for 2 h at 4 °C. Decant the supernatant liquid into a clean tube (3.3.4) and dilute with
400 ml of ultrapure water prior to filtration.
4.2.2.3 Purification of polymer particles by ultracentrifugation
3 3 3 3
Place 2 ml of each sucrose solution (3.2.13) (ρ = 1,35 g/cm , 1,17 g/cm , 1,08 g/cm , 1,04 g/cm and
3
1,02 g/cm ) into centrifuge tubes (3.3.4) so that the tubes are roughly three-quarters full, and apply measured
aliquots of the digested tissue suspension to the surface of the sucrose solution in each tube. Ultracentrifuge
at 100 000 g for 3 h at 5 °C. Carefully collect the top layer into a sterile tube and dilute with distilled water at
65 °C to help dilute the residual sucrose. Ultrasonicate for 10 min to break up the agglomerated particles and
then heat for 1 h at 80 °C to dissolve the sucrose.
Apply measured volumes of the suspension to two layers of isopropanol-water mixture (3.2.6) of densities
3 3
0,90 g/cm and 0,96 g/cm formed in ultracentrifuge tubes. Ultracentrifuge these at 100 000 g for 1 h at 20 °C.
After removing the tubes from the ultracentrifuge rotor, a layer of white particles should be visible at the
interface of the two layers. Remove this layer, containing the polyethylene particles, and place into a sterile
tube using a fine-tipped glass pipette (3.3.12) inserted through the top isopropanol layer. Ultrasonicate for
10 min to break up any aggregates.
Different ultracentrifugation times and speeds may be used, provided that they have been demonstrated to
give the same degree of separation and the results of the verification procedure have been documented.
NOTE 1 The first ultracentrifugation step serves to separate the lighter polyethylene wear particles from the heavier
fractions. The second ultracentrifugation step purifies the polyethylene particle yield by putting it through a finer density
gradient.
NOTE 2 This method might discriminate against the largest sizes of polyethylene generated, and consequently the
total wear volume might not be isolated.
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ISO 17853:2010(E)
4.3 Procedure for metal particle isolation
Due to the solubility of metals in strong acids and alkalis, an enzymatic digestion method needs to be used.
[5]
The method below has been described by Catelas et al. and is similar to the procedure developed earlier by
the same authors for particle isolation from joint simulator lubricant {see Reference [6] (c.f. Clause 5)}, with
only minor differences in the initial steps as well as in the enzyme concentrations to account for the use of
tissue instead of serum lubricant.
NOTE 1 Being able to use the same procedure to isolate and characterize particles from tissues and joint simulator
lubricant enables direct and accurate comparison of the isolated particles. This constitutes a significant advantage to this
procedure.
a) Cut the tissue into small pieces using a scalpel and blade to speed up the digestion time. Resuspend
several small pieces (about 2 mm × 2 mm × 2 mm) in 2 ml tubes.
NOTE 2 The tissue weight will depend on the overall wear noticed in the patient as well as the piece of tissue used for
particle isolation (e.g. granuloma, capsule), but a minimum of 200 mg wet weight is recommended.
b) Wash four times for 2 min in sodium phosphate buffer (3.2.8), pH 7,4.
c) Resuspend the tissue pieces in 1 ml of sodium dodecyl sulfate (3.2.11) (2,5 g/100 ml solution in distilled
water) and boil for 10 min. While boiling, homogenize the tissue pieces in solution using a Teflon-glass
potter tissue grinder (3.3.18) every 2 min.
d) Cool at room temperature for 10 min.
e) Centrifuge the tubes at 16 000 g for 10 min.
f) Wash once with 1 ml of acetone (3.2.2), diluted with distilled water with a volume fraction of 80 % acetone.
Centrifuge at 16 000 g for 10 min.
g) Wash three times with 1 ml of 250 mM sodium phosphate buffer containing 25 mM EDTA, pH 7,4.
Centrifuge at 16 000 g for 10 min for each wash.
h) Sonicate in 1 ml of 250 mM sodium phosphate buffer containing 25 mM EDTA, pH 7,4, for 20 s to 25 s,
using an ultrasonic cell disrupter equipped with a microprobe, or in a sonicating water bath for 30 min.
NOTE 3 Using the ultrasonic cell disrupter is more efficient, but an appropriate apparatus with a clean and
undamaged/non-corroded probe tip is used to avoid potential titanium contamination from the probe tip.
i) Add 0,5 ml of 250 mM sodium phosphate buffer containing 25 mM EDTA, pH 7,4, and papain (3.2.7)
(4,8 units per 1,5 ml of phosphate buffer). Incubate in an agitated water bath (3.3.22) for 24 h at 65 °C.
j) Centrifuge the tubes at 16 000 g for 10 min.
k) Carefully remove the liquid using a micropipette without touching the pellet at the bottom of the tubes.
l) Resuspend the pellet in 1 ml of sodium dodecyl sulfate (2,5 g/100 ml solution in distilled water).
m) Boil for 10 min.
n) Cool at room temperature for 10 min.
o) Centrifuge the tubes at 16 000 g for 10 min.
p) Carefully remove the liquid using a micropipette without touching the pellet at the bottom of the tubes.
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ISO 17853:2010(E)
q) Wash the pellet with 1 ml of 50 mM TRIS-HCl (3.2.14) (pH 7,6). Centrifuge the tubes at 16 000 g for
10 min. Carefully remove the liquid using a micropipette without touching the pellet at the bottom of the
tubes.
r) Repeat step q) once more.
s) Resuspend the pellet in 1 ml of 50 mM TRIS-HCl and sonicate for up to 30 s using an ultrasonic cell
disrupter equipped with a microprobe, or in a sonicating water bath for 30 min.
NOTE 4 Once again, using the ultrasonic cell disrupter is more efficient, but an appropriate apparatus with a clean and
undamaged/non-corroded probe tip is used to avoid potential titanium contamination from the probe tip.
t) Add proteinase K (3.2.10) (2 g/ml of TRIS-HCl buffer) and incubate for 24 h at 55 °C in an agitated water
bath.
u) Centrifuge the tubes at 16 000 g for 15 min.
v) Carefully remove the liquid using a micropipette without touching the pellet at the bottom of the tubes.
w) Add 1 ml of sodium dodecyl sulfate (2,5 g/100 ml solution in distilled water).
x) Boil for 10 min.
y) Cool at room temperature for 10 min.
z) Centrifuge the tubes at 16 000 g for 15 min.
aa) Carefully remove the liquid using a micropipette without touching the pellet at the bottom of the tubes.
bb) Wash with 1 ml of 50 mM TRIS-HCl (pH 7,6). Centrifuge at 16 000 g for 15 min and carefully remove the
liquid using a micropipette without touching the pellet at the bottom of the tubes.
cc) Wash with 0,5 ml of acetone prepared in 3 % SDS (3.2.11), with a volume fraction of 80 % acetone.
Centrifuge at 16 000 g for 15 min and carefully remove the liquid using a micropipette without touching
the pellet at the bottom of the tubes.
dd) Wash with 1 ml of distilled water. Centrifuge at 16 000 g for 15 min and carefully remove the liquid using a
micropipette without touching the pellet at the bottom of the tubes.
ee) Add absolute ethanol (3.2.1) and store at 4 °C until collecting the particles (see 4.4.2).
NOTE 5 Alternatively, steps g) to i) might be performed using 50 mM TRIS-HCl buffer. The efficacy of the papain
enzyme used should then be verified.
4.4 Collection of particles
4.4.1 Polyethylene particles
Collect particles on filters of pore size 0,1 µm followed by a 0,015 µm filter, or similar. For particles isolated by
high-speed centrifugation or where the whole wear volume is necessary, filter the entire sample volume
obtained in 4.2.2.2 using a vacuum filtration system. For particles isolated by ultracentrifugation (see 4.2.2.3),
use between 10 µl and 300 µl aliquots of the particle suspension or larger volumes of diluted suspension.
NOTE The appropriate dilution is achieved when the suspension appears almost clear, usually at dilutions between
1:10 and 1:100. The aim is to produce a concentration of particles on the filter which is not so dense as to make
visualization of discrete particles difficult by SEM, while ensuring a reasonable number of particles is available for analysis
(at least 100 particles).
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ISO 17853:2010(E)
If diluting, record the exact dilution for each sample. For filtration, take up a known volume of the particle
suspension into a syringe (3.3.17) and attach the end of the syringe to the filtration unit (3.3.7). Apply gentle
pressure to the syringe in order to push the water through the filter and out at the other end of the filtration unit
at a rate of about 1 drop (0,045 ml) per second, depositing the particles on the surface of the filter. Change the
filter if it becomes blocked. Use the following procedure on each filter. Flush the filter and syringe through with
distilled water. Finally, flush the filter with air in the same direction as the filtration before removing the filter
from the unit, using tweezers, and leaving it to dry in a sterile, covered Petri dish.
Due to the small size of the particles, filters with a pore size smaller than 0,1 µm should be used, if available.
In this case, sequential filtering might be necessary to avoid pore clogging.
4.4.2 Metal particles
Centrifuge particles in ethanol (from the isolation protocol described in 4.3) at 16 000 g for 20 min. Remove
supernatant liquid and add 0,5 ml of 100 % acetone:epoxy resin (1:1). Place the tubes in a tube holder and
rotate overnight at room temperature (this step could possibly be shortened for very small pellets). Centrifuge
the particles at 16 000 g for 20 min. Carefully remove the liquid using a micropipette and without touching the
pellet at the bottom of the tube. Leave the tubes under vacuum for 1 h to remove the remaining acetone. Add
1 ml of pure epoxy resin (3.2.10) and leave the tubes under vacuum for 3 h. Place the tubes at 60 °C for 48 h
to allow resin polymerization. Remove plastic tubes to obtain solid resin with the particle pellets at the bottom.
This protocol will enable resin infiltration and particle separation.
Section pellets using a diamond knife and spread the sections (about 90 nm to 120 nm thick) on
formvar-coated copper grids of 200 mesh size for TEM analysis. See References [5] and [6]. Section the
pellets such that an even representation of all the particles in the pellets can be ascertained (section the whole
pellet if very small).
NOTE Metal particles can also be collected by vacuum filtration on a 0,015 µm filter membrane for analysis using
high-resolution SEM; see Reference [7]. However, when using this method, the user needs to check for potential particle
agglomeration and oxidation on the filter and be aware of potential particle loss.
4.5 Particle size and shape characterization
4.5.1 Polyethylene particles
For SEM imaging of particles, attach the filter with particles on it to an SEM mount using a carbon sticker
(3.3.3). Coat the filter with gold or other conductive material, e.g. platinum/palladium, to make the particles
conductive. The coating thickness shall be 3 nm to 5 nm. Image the polymer particles at an accelerating
voltage of not greater than 10 keV.
To characterize particles larger than 0,1 µm, select random, non-overlapping fields on the filter carrying the
particles at a magnification of at least ×5 000 until a total of 100 particles has been imaged. For particles
larger than 10 µm, use a lower magnification such as ×500.
Characterize the size, shape and area of the particles using a series of predefined descriptions such as length,
breadth, equivalent circle diameter (diameter of a circle with the same area as particle), area, perimeter,
[8]
aspect ratio (length:breadth) and roundness (perimeter 1/4π × area), as described in ASTM F 1877-05 . The
magnification at which size and shape analysis was performed should be stated in the test report.
To characterize particles smaller than 0,1 µm, use high-resolution SEM. Use magnifications up to ×100 000 at
an accelerating voltage of 3 keV.
NOTE 1 The differentiation of fibrillar and rounded polymer particles can also be useful.
NOTE 2 Computerized or manual image analysis software can be used to determine particle sizes and shapes.
NOTE 3 Particle analysers might also be used if their limit of resolution is 0,1 µm or less, however, there is a risk of
size overestimation due to particle agglomeration.
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4.5.2 Metal particles
4.5.2.1 General
As described in 4.4.2, particles are typically analysed by TEM, but they could also be analysed by SEM.
4.5.2.2 TEM analysis
Image metal particles using TEM at an accelerating voltage of around 80 kV and a magnification setting of at
least ×21 000 (the magnification should be stated in the report).
Characterize metal particles in terms of size and shape by manual image analysis of representative TEM
micrographs (minimum 150 particles, in case of low wear; 300 or more, if permitted). Characterize all particles
from each micrograph in order to have a better and unbiased representation of the different particle sizes.
Determine the maximum dimension (or length) of each part
...
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