1. health and fitness
2. disease

Biofilm Formation in Milk Production and
Processing Environments; Influence on Milk
Quality and Safety
Sophie Marchand, Jan De Block, Valerie De Jonghe, An Coorevits, Marc Heyndrickx, and Lieve Herman
Abstract: Bacteria in milk have the ability to adhere and aggregate on stainless steel surfaces, resulting in biofilm
formation in milk storage tanks and milk process lines. Growth of biofilms in milk processing environments leads to
increased opportunity for microbial contamination of the processed dairy products. These biofilms may contain spoilage
and pathogenic microorganisms. Bacteria within biofilms are protected from sanitizers due to multispecies cooperation
and the presence of extracellular polymeric substances, by which their survival and subsequent contamination of processed
milk products is promoted. This paper reviews the most critical factors in biofilm formation, with special attention to
pseudomonads, the predominant spoilage bacteria originating from raw milk. Biofilm interactions between pseudomonads
and milk pathogens are also addressed, as emerging risks and future research perspectives, specifically related to the milk
processing environment.
Introduction
Raw milk is an ideal culture medium for microorganisms. Because the microbial load of milk may hold spoilage and/or health
risks, the manufacture of milk and milk products is subject to very
stringent rules. These rules cover the way in which livestock is
kept and milked, milk storage facilities, preparation methods, additives, processing equipment, and the transport tanks that move
milk from the farm to the processing plants (Anonymous 2005;
Anonymous 2006; Anonymous 2007; Anonymous 2011). On its
journey from the farm to the consumer, milk comes into contact
with the walls of the equipment in which it is being processed and
transported. Since the European (and American) legislation has
strict regulations concerning materials coming into contact with
foods (Anonymous 2004; FDA 2007; EFSA 2008) and milk processing necessitates hygienic equipment material resistant to corrosion in alkaline and/or acidic conditions (Boulangé-Petermann
and others 1997), the dairy industry has employed stainless steel for
more than 60 years in almost all segments of the dairy chain. The
development of stainless steel in the dairy industry is explained by
the fact that it corresponds exactly to the requirements expected of
materials in contact with food: 1) the material has to be chemical,
MS 20110997 Submitted 8/18/2011, Accepted 12/13/2011. Authors Marchand,
De Block, De Jonghe, Heyndrickx, and Herman are with the Inst. for Agricultural and Fisheries Research—Technology and Food Sciences Unit (ILVO-T&V),
Brusselsesteenweg 370, 9090 Melle, Belgium. Author Coorevits is with the Faculty
of Applied Engineering Sciences, Dept. of Biochemistry and Brewing, Univ. College
Ghent, Schoonmeersstraat 52, 9000 Gent, Belgium; and Faculty of Science, Dept.
of Biochemistry and Microbiology, Laboratory of Microbiology, Ghent Univ., K. L.
Ledeganckstraat 35, 9000 Ghent, Belgium. Direct inquiries to author Marchand
(E-mail: [email protected]).
c 2012 Institute of Food Technologists®
doi: 10.1111/j.1541-4337.2011.00183.x
bacteriological, and organoleptical neutral with regard to the food
product, 2) the material should be easy to clean so that the hygiene
and appearance of the food product are guaranteed, and 3) it has
to be durable, including corrosion and aging (Anonymous 2004;
Bremer and others 2009). Other factors also contribute to the
preference of the dairy industry for stainless steel. These include
its mechanical characteristics, expansion coefficient, thermal conductivity, and ease of use (Bremer and others 2009). It is difficult to
find alternative products to compete with stainless steel in the milk
industry, because of the processing conditions. However, in some
manufacturing operations, alternative materials can be employed,
but their use is still limited and restricted to certain applications.
Examples of nonmetal materials used are elastomers (also known as
rubbers) and plastics. They are often used in conveyer belts, containers, seals, gaskets, or cutting boards. Rubbers, such as ethylene
propylene diene monomer rubber (EPDM), nitril butyl rubber
(NBR, known as Buna-N® ), silicon rubber, or fluoroelastomer
(Viton) are used in both closed equipment (seals gaskets, membranes, fittings, and containers) and in open equipment such as
conveyer belts (Faille and Carpentier 2009). Among these materials, the most frequently used gasket materials in milk processing
equipment are EPDM and NBR (Faille and Carpentier 2009). A
wide range of plastics is also available, but only a few of them are
food-approved, such as polypropylene (PP), polycarbonate (PC),
high-density polyethylene (HDPE), unplasticized polyvinyl chloride (PVC), and fluoropolymers such as polytetrafluoroethylene
(PTFE, Teflon® ). The latter, used for gaskets in the food industry,
is porous and lacks resilience and must thus be used with care
(Faille and Carpentier 2009).
Surfaces of equipment used in food and beverage (such as
milk) processing and handling are commonly contaminated by
Vol. 11, 2012 r Comprehensive Reviews in Food Science and Food Safety 133
Biofilm risks in dairy practice . . .
Figure 1–Stages of bacterial biofilm development. Adapted from Stoodley and others (2002). Stage 1: Initial attachment of cells to the surface.
Stage 2: Production of EPS resulting in more firmly adhered “irreversible” attachment. Stage 3: Early development of biofilm architecture.
Stage 4: Maturation of biofilm architecture. Stage 5: Dispersion of single cells from the biofilm. The bottom panels show each of the 5 stages of
development represented by a photomicrograph of P. aeruginosa when grown under continuous-flow conditions on a glass substratum.
microorganisms, even following cleaning and disinfection procedures (Gibson and others 1999; Marouani-Gadri and others 2010).
These contaminating microorganisms appear as adherent microorganisms or as more complex structures called biofilms. Adherent
spores and bacteria, as well as biofilms, can be observed on every
surface of food industry plants such as stainless steel surfaces (Figure 2), floors, belts, or rubber seals (Costerton and others 1995;
Kumar and Anand 1998).
Understanding Biofilms
An important reservoir of microbial contamination that has received relatively little attention in the dairy industry is the microbial biofilm. In milk storage and dairy processing operations,
as well as in numerous other industrial systems, besides being
present in the raw material, most bacteria are associated with surfaces (Mittelman and others 1990; Mosteller and Bishop 1993;
Mittelman 1998). The attachment of “pioneering” bacteria with
subsequent development of biofilms in milk processing environments is a potential source of contamination of finished products
that may shorten the shelf life or facilitate transmission of diseases (Hood and Zottola 1995; Lindsay and others 2002; Brooks
and Flint 2008). Despite the fact that bacteria are predominantly
present in biofilms, for many years studies on bacterial physiology
have focused primarily on the planktonic state. Now, however, it
is well established that bacteria are able to switch between different habitation modes: single cells (the planktonic or free floating
state) and biofilms. In addition, it has been established that for each
planktonic bacterium detected, there might be close to 1000 or-
ganisms present in biofilms (Momba and others 2000). A biofilm is
defined as a sessile microbial community characterized by adhesion
to a solid surface and by production of a matrix that surrounds the
bacterial cells and includes extracellular polysaccharides (EPSs),
proteins and DNA (Wingender and others 2001; Whitchurch and
others 2002; Costerton and others 2003; Bjarnsholt and others
2009). Biofilm development is a result of successful attachment
and subsequent growth of microorganisms on a surface (Figure 1).
Under suitable conditions, a biofilm in a milk processing environment develops initially through accumulation of organic matter
on a metal surface, which is then colonized by bacteria. Transition
from planktonic mode to biofilm mode is regulated by a variety of
environmental and physiological triggers, such as quorum sensing,
nutrient availability, and cellular stress. A biofilm community may
comprise single and/or multiple species of bacteria and form a single layer or 3-dimensional structures. Biofilms are large, complex,
and organized bacterial ecosystems in which water channels are
dispersed providing passages for nutrient, metabolite, and waste
product exchange (Sauer and others 2007). Biofilm communities
can even provide (in analogy with apoptosis in higher eukaryotes)
the selective pressure that is required for programmed cell death,
by eliminating damaged individuals from the population (Bayles
2007). Because of competition reduction and the release of nutrients from the dead and lysed cells, nutrient availability is more
easily maintained for the healthy individuals that remain (Bayles
2007). Programmed death and lysis of the bacterial cells probably
occurs as a function of their spatial orientation within the biofilm.
In addition, the released genomic DNA is a structural component
134 Comprehensive Reviews in Food Science and Food Safety r Vol. 11, 2012
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Biofilm risks in dairy practice . . .
Figure 2–Adhesion of various microorganisms on stainless steel surfaces. Adapted from Faille and Carpentier (2009). (A–B) Adhesion of
Staphylococcus caprea and Pseudomonas fluorescens under static condition on stainless steel with a 2B finish (horizontally immersed). (C–D) Adhesion
of Bacillus cereus spores on substrata with irregular topography, vertically immersed in a spore suspension (presence of scratches, flaws, . . . ).
of the biofilm matrix, which supports the notion that cell lysis
contributes to the stability of the overall biofilm structure (Bayles
2007).
According to Mittelman (1998), the making of a mature biofilm
may take several hours to several weeks, depending on the system
under development. For example, in an experiment with Pseudomonas aeruginosa, a common biofilm former on medical devices,
it has been established that attachment to stainless steel took place
within 30 s of exposure; in an industrial water simulation experiment in a biofilm annular reactor, the colony forming units
(CFUs) within the biofilm increased approximately 5-fold, from
420 to 2123 CFU/15 cm2 , as the incubation time was prolonged
from 24 to 96 h (Florjanic and Kristl 2011). More importantly,
in dairy equipment biofilms, the development is also very rapid
(8–12 h) (Scott and others 2007; Bremer and others 2009), with
numbers of up to 106 bacteria per cm2 being recorded in the generation section of a pasteurizer after 12 h of operation (Bouman and
others 1982; Bremer and others 2009). While a biofilm can spread
at its own rate by ordinary cell division, it will also periodically
release “pioneer” cells to colonize downstream sections of piping.
The biological, chemical, and physiological factors that drive detachment are complex and incompletely understood (Chambless
and Stewart 2007). Multiple factors are probably associated with
attachment and detachment processes, depending on the availability of nutrients or oxygen (Chandy and Angles 2001; Rice
and others 2005), shear–stress (Mittelman 1998; Guillemot and
others 2006; Lee and others 2008; Florjanic and Kristl 2011),
quorum sensing (Rice and others 2005), microbial metabolic activity, and microbial gene expression (Kaplan and others 2003;
Kaplan and others 2004). Biofilm detachment has been divided
c 2012 Institute of Food Technologists®
into 3 processes: erosion, abrasion, and sloughing (Garny and others 2008). Erosion (result of fluid shear forces) and abrasion (collision of particles) refer to the continuous detachment of single cells
or small cell clusters and affect the total biofilm surface. Sloughing refers to the instant loss of large parts of the biofilm, therefore affecting the entire biofilm and not only the biofilm surface
(Morgenroth 2003). Depending on the strength of the biofilm,
sloughing can even lead to a complete loss of the biofilm. Several
detachment processes may occur simultaneously (Telgmann and
others 2004). However, the original biofilm structure and magnitude and the detachment force might have a strong influence on
the frequency and extent of a specific detachment process.
Biofilms are characterized by environmental conditions and the
surfaces colonized, the bacterial genes activated and required to
form and maintain the biofilm, and the types of extracellular products that are concentrated in the biofilm matrix. There are many
different types of biofilms and even one bacterium may make
several different types of biofilms under different environmental
conditions. Here, we review the diverse array of biofilms formed
in milk processing environment, with special attention to pseudomonads, the predominant spoilage bacteria originating from
raw milk. Biofilm interactions between pseudomonads and milk
pathogens will also be addressed, as well as emerging risks and
biofilm control strategies specifically related to the milk processing environment.
Dairy Practice—Mechanisms of Biofilm Formation
Bacterial attachment and the formation of biofilms appear to
take place in different stages, such as formation of a conditioning layer, bacterial adhesion, bacterial growth, and biofilm
Vol. 11, 2012 r Comprehensive Reviews in Food Science and Food Safety 135
Biofilm risks in dairy practice . . .
expansion (Kumar and Anand 1998; O’Toole and others 2000;
McLandsborough and others 2006; Kokare and others 2009). In
dairy operations, the conditioning film mainly consists of organic milk components. This first stage occurs within the first
5–10 s after placement of an otherwise clean surface into a fluid
environment (Mittelman 1998). The conditioning also alters the
physicochemical properties of the surface, such as surface free energy, changes in hydrophobicity, and electrostatic charges, which
may affect the subsequent order of microbial events (Dickson and
Koohmaraie 1989). The formation of conditioning films can be
influenced by the material type contacting the milk. As an example, certain materials are known for their “theta surface,” which
is a characteristic expression of outermost atomic features least
retentive of depositing proteins, and identified by the bioengineering criterion of having a measured critical surface tension
(CST) between 20 and 30 mN/m (Baier 2006). The most effective atomic group exposures for theta surface results are intrinsically hydrophobic, closely packed methyl, CH3 , terminals, or
repeating CH2 CF2 runs in polyvinylidene fluoride (PVDF) (Baier
2006). Unfortunately, most of these materials are not (yet) approved, at least in Europe, for contact with food (Anonymous
2004). Therefore, further research is needed before applications
in the food industry become possible. During the second stage
of biofilm formation, single bacterial cells are transported to surfaces and reversible bonds are formed between the cell wall and
the substratum. Bacterial attachment is mediated by fimbriae, pili,
flagella, and bacterial extracellular polymeric substances (EPSs)
that act to form a bridge between bacteria and the conditioning
film (Kokare and others 2009). The chemical structure of the EPS
varies among different types of organisms and is also dependent
on environmental conditions (Momba and others 2000). While
there is some debate about the influence on surface roughness
on bacterial attachment (Sreekumari and others 2005; Oliveira
and others 2006; Silva and others 2008), there appears to be a
general agreement about the importance of using surfaces with
minimal cracks and crevices in order to reduce bacterial adherence and biofilm growth and to enhance cleaning effectiveness (Bremer and others 2009). Once established, biofilms accelerate corrosion and material detoriation (Storgards and others
1999a). Dead ends, corners, cracks, crevices, gaskets, valves, and
joints are all possible points for biofilm formation (Storgards and
others 1999a; Storgards and others 1999b). Biofilms do not possess a uniform structure (Wimpenny and others 2000; McLandsborough and others 2006). The structures that are formed depend on a large variety of intrinsic and extrinsic factors such as
species, temperature, flow conditions, pH, presence of salts, nutrients, and so on (McLandsborough and others 2006). Next to contact material, temperature plays an important role in the adhesion
of bacteria to surfaces. In general, higher temperatures (37 ◦ C in
comparison with 4, 12, and 22 ◦ C) seem to increase cell surface hydrophobicity and subsequently bacterial attachment (Cappello and
Guglielmino 2006; Di Bonaventura and others 2008). The effect
of flow conditions has not been well studied in the dairy industry,
but from studies in other systems, it is known that biofilms grow
denser under high than under low shear conditions (Stoodley and
others 2002; Bremer and others 2009). Contrary to expectations,
both laminar and turbulent flow conditions have been observed
to enhance bacterial attachment (by bringing bacteria closer to a
surface) when compared to static conditions (Rijnaarts and others
1993). It has been speculated that turbulent flow may push bacterial cells onto the surface, thus enhancing probability of adhesion
and biofilm formation (Donlan and Costerton 2002). With regard
to pH, it has been shown that the pH of the surrounding solution
influences the interaction between bacterial cells and the metal
surface; the bacteria–metal adhesion force appears to reach the
highest value when the pH of the solution is near the isolelectric
point of the bacteria, that is, at the zero point charge (Sheng and
others 2008). Stronger ionic strength in the solution, on the other
hand, results in a higher bacteria–metal adhesion force, which
is due to the stronger electrostatic attraction force between the
positively charged metal surface and negatively charged bacterial
surface (Sheng and others 2008). From this, it can be deduced
that the higher the adhesion force, the more bacteria will attach
to a particular surface. Concerning the effect of the presence of
nutrients or certain milk components on the adhesion of bacteria,
some conflicting statements can be retrieved from the literature.
On the one hand, it is stated that milk proteins coated on stainless steel, rubber, and dairy equipment reduce bacterial adhesion
(Speers and Gilmour 1985; Helke and others 1993; Bernbom and
others 2009), while, on the other hand, certain bacteria (for example, Bacillus cereus) appear to need certain milk components before
adhesion can occur (Shaheen and others 2010). These contradictory findings might be explained by the fact that different milk
types (skim compared with whole milk and heated (100 min for
30 min) compared with unheated) were used in the experimental
setups. Evidently, denatured milk proteins may have other characteristics than undenatured proteins naturally present in refrigerated
whole raw milk. Second, whole milk contains natural surfactants
and phospholipids, both surface-active compounds that can be retrieved in the fat globules of milk. The ability of B. cereus spores
to adhere and act as an initiation stage for biofilm formation on
a wide variety of materials commonly encountered in food processing plants is also well known (Peng and others 2001; Faille and
others 2001; Heyndrickx and others 2010). The strong adhesion
properties of B. cereus spores have been attributed to the hydrophobic character of the exosporium (Peng and others 2001; Faille and
others 2001), which varies from species to species (Tauveron and
others 2006) and to the presence of appendages on the surface
of the spores (Vanloosdrecht and others 1989). Thick biofilms
of B. cereus were shown to develop on stainless steel coupons at
the air–liquid interface, while biofilm formation was much lower
in submerged systems (Wijman and others 2007). This suggests
that B. cereus biofilms develop particularly in partly filled industrial storage and piping systems and these biofilms act as a shelter
for spore formation that can be subsequently released by dispersal
into the food production system. Spores embedded in biofilms are
protected against disinfectants such as chlorine, chlorine dioxide,
and peroxyacetic-acid-based sanitizer (Ryu and Beuchat 2005). In
some dairies, persistent silo tank contamination, heat exchange
equipment contamination, or postpasteurization contamination
are important sources of B. cereus (te Giffel and others 1996a;
te Giffel and others 1996b). For pasteurized and extended shelf
life (ESL) milk, the filling machine has been shown as the main
source of recontamination, with the filler nozzles, aerosols, and
the water at the bottom of the filling machine being of particular
concern (Rysstad and Kolstad 2006). While for most of Bacillus
strains, negative effects of whole milk on biofilm formation have
been observed (Flint and others 1997a; Wong 1998), the study of
Shaheen and others (2010) illustrated that B. cereus was capable of
forming biofilms in whole milk, but not in water-diluted milk.
The results of that latter study suggest that any surface-active compound found in whole milk might work as a surfactant needed
for biofilm formation by certain strains of B. cereus. However, the
effects of surfactants on biofilm formation might be strain-specific
136 Comprehensive Reviews in Food Science and Food Safety r Vol. 11, 2012
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Biofilm risks in dairy practice . . .
and generalization of results concerning (anti-)adhesive properties
of milk compounds should thus be avoided and evaluated carefully.
Several groups have reported on the ability of bacteria to attach
to surfaces commonly found in the milk processing environment,
such as rubber and stainless steel (Czechowski 1990; Krysinski and
others 1992; Suarez and others 1992). Scanning electron micrographs revealed that food-borne pathogens and spoilage microorganisms can accumulate as biofilms on aluminum, Buna-N and
Teflon seals, and nylon materials typically found in food processing environments (Herald and Zottola 1988a; Herald and Zottola
1988b; Mafu and others 1990; Blackman and Frank 1996). More
importantly, during heat processes above 65 ◦ C, whey proteins in
milk begin to denature and aggregate, which can lead to a faster
adherence than proteins in their native state. This protein adherence can change the surface properties of stainless steel (De Jong
1997), increasing the likelihood of bacteria attaching to a surface
and creating an environment that encourages bacterial attachment
to an extent where in one study, it was discovered that fouled surfaces attracted 10–100 times more vegetative cells and spores of G.
stearothermophilus to the surface than the clean stainless steel (Flint
and others 2001). The milking equipment can be contaminated by
milk spoilers and pathogens through the dairy farm and processing environment, but also through the rinsing water used in the
milking machines (Oliver and others 2005). Microorganisms originating from rinsing water (especially Pseudomonas, Aeromonas, and
Legionella spp. [Momba and others 2000]) can form biofilms that
are difficult to eradicate and can act as a harbor and/or substrate
for other microorganisms less prone to biofilm formation, thus
increasing the probability of pathogen survival and further dissemination during milk processing (Lomander and others 2004).
Table 1 demonstrates typical problem areas within dairies. Teixeira
and others (2005) also illustrated that the short rubber milking
tube (of the cluster in automatic milking machines) is one of the
points more prone to biofilm formation. The cluster, which attaches to the udder of the cow, consists of 4 teatcup assemblies
(each having a shell, a rubber liner, and a short milk and short
pulse tube), a claw, a long milk tube, and a long pulse tube. All
these constituents are made of rubber, stainless steel, or plastic.
Other possible hazards include biofilm accumulation and microbial colonization in milk pipelines, storage tanks, and milk silos
(Shaheen and others 2010), as well as fouling of heat exchangers
(Giffel and others 1997; Flint and others 1997b; Flint and others
1999; Flint and others 2000) and adhesion of spores on packaging
material surfaces (Kirtley and Mcguire 1989).
Environments, which select for monospecies biofilms (such
as those of thermophilic bacilli) in dairy processing plants, are
typically the sections with elevated temperatures (40 to 65 ◦ C)
(Stadhouders and others 1982; Flint and others 1997a; Murphy
and others 1999). Examples are preheating and evaporation sections of milk powder plants, plate heat exchangers used during
the pasteurization process, centrifugal separators operated at warm
temperatures (45–55 ◦ C), recycle loops in butter manufacturing
plants, and cream heaters in anhydrous milk fat plants (Burgess and
others 2010). A typical problem in the manufacture of milk powder is the high levels of Anoxybacillus flavithermus and Geobacillus
spp. The spores of these organisms are very heat-resistant, with the
vegetative cells able to grow in temperature of up to 65 ◦ C (Palmer
and others 2010). The bacteria are normally present in low levels
in raw milk, but may reach 105 CFU/g in the final product after
15–20 h of plant operation (Hinton and others 2002; Ruckert and
others 2004). The limited residence time of the milk during milk
powder manufacture cannot explain the number of thermophiles
c 2012 Institute of Food Technologists®
found in the final product. Suggestions are made that biofilm formation on the milk evaporator and consequent sloughing off into
the product line is responsible for the high contamination levels
of the final product (Hinton and others 2002; Palmer and others 2010). Strategies (such as shorter production lengths and the
use of sanitizers) to prevent thermophilic biofilm formation had
limited success, partly due to limited knowledge on the structure
and composition of those biofilms in milk processing operations
(Burgess and others 2010).
In a model pasteurizer, thermophilic streptococci were detected
on the walls of the cooling section at levels of 107 cells/cm2 ,
and subsequent research in processing facilities indicated that
thermophilic streptococci could be frequently isolated from the
cooling section of pasteurizers (Bouman and others 1982). The
attachment of resistant Streptoccocus thermophilus occurs mainly to
heat exchanger plates in the downstream side sections of pasteurizers giving rise to the contamination of pasteurized milk (Driessen
and others 1984). Recontamination of consumer packages of pasteurized milk with Gram-negative psychrotrophic bacteria, on the
other hand, was associated with rinsing water in and around the
filling machine during the filling operation (Eneroth and others
1998; Dogan and Boor 2003). This suggests that bacteria could
have formed biofilms in the rinse water system. Biofilms that
can develop on the sides of gaskets may also be a possible source
of postpasteurization contamination (Austin and Bergeron 1995).
Langeveld and others (1995) heated milk in a laboratory-scale
stainless steel tube heat exchanger and found that, as a result of
release from the tube walls, the concentration of bacteria in the
milk could increase by a factor of 106 . These authors demonstrated a relationship between the density of bacteria on the tube
walls and the concentration of cells in the milk after heating.
The observations of Scott and others (2007) on commercial milk
evaporators showed that in a plant cleaned according to standard industrial practice and processing high-quality milk (<100
thermophiles per mL) that resides, on average, 20–30 min in the
plant, might result in outflowing milk that can contain up to
106 cells per mL within 18 h of run commencement. The authors
concluded that it was not possible for these numbers to have been
produced during the transit of the milk through the plant and must
thus have originated from cells immobilized on plant internal surfaces. Other locations where biofilms often arise are ultrafiltration
and reverse osmosis membranes (Tang and others 2009a, 2009b,
2010). Membrane separation technology is often used for the removal of bacteria from skim milk in the production of ESL milk,
concentration of casein micelles, and recovery of serum proteins
from whey. Membrane biofouling caused by microbial attachment
leads to decreased membrane flux and increased filtration pressure, and subsequently, increased operation cost due to frequent
cleaning and replacement of clogged membranes (Liao and others
2004; Le-Clech and others 2006). Tang and others (2009a) illustrated that the bacterial isolates recovered from such membranes
in dairy plants predominantly belonged to Pseudomonas, Bacillus,
and Klebsiella genera. Furthermore, the attachment of the different
isolates appeared highly variable and there was an enhanced adherence in the presence of whey. In an ice cream plant, most of the
biofilm formations were seen on the conveyer belt of the packaging machine 8 h after the beginning of the production (Gunduz
and Tuncel 2006). Most of the Gram-negative biofilm-forming
bacteria were identified as Proteus, Enterobacter, Citrobacter, Shigella,
Escherichia, Edwardsiella, Aeromonas, Plesiomonas, Moraxella, Alcaligenes, and Pseudomonas species. Gram-positive biofilm-forming
isolates consisted of Staphylococcus, Bacillus, Listeria, and lactic acid
Vol. 11, 2012 r Comprehensive Reviews in Food Science and Food Safety 137
Biofilm risks in dairy practice . . .
Table 1–Overview of biofilm problem areas at dairy farms and dairy processing plants (Wirtanen 2004; Teixeira and others 2005; Agarwal and others
2006; Gunduz and Tuncel 2006).
Sampling points
Balance tank
Aging tank
Feeding unit
Conveyer belt of packaging machine
Floor drain
Doormat
Ultrafiltration membranes
Silo, welded joints
Valves
Air separators, inside
Tank truck, valve, gasket
Tank truck, air separator
Tank truck, air separator, gasket
Bulk tank outlet on farm
Rubber liners
Short milking tube
Materials
Steel
Steel
∗
Rubber
∗
∗
Steel
Steel
Steel
Steel
Rubber
Steel
Rubber
Steel
Rubber
Rubber
Pseudomonas
−
−
−
+
−
−
−
+
−
+
+
+
−
∗
∗
+
Aeromonas
−
−
−
−
−
+
−
−
−
−
−
−
−
−
−
−
bacteria such as Streptococcus, Leuconostoc, and Pediococcus (Gunduz
and Tuncel 2006) species. A brief overview of bacteria isolated
at different sampling points in dairy processing plants is given in
Table 1.
Which organisms attach to the surface is a function of the planktonic population present in the raw material and the processing
conditions in the particular equipment. Heat-sensitive Pseudomonas
and Listeria species are most likely to be found in pipes and silos holding milk prior to pasteurization, whereas thermophilic
biofilms may form in heated equipment. In terms of their effect
on product acceptability, biofilms can contain a dual risk: product
detoriation and disease transmission, respectively, through spoilage
bacteria and pathogens.
Risks posed by spoilage bacteria and pathogens—the role
of pseudomonads
Bacterial spoilage still causes significant losses for the dairy
industry. Milk contamination with psychrotrophic microorganisms is of particular concern to the dairy industry as dairy products are stored and distributed at temperatures permissive for the
growth of these organisms. Psychrotrophic bacteria are ubiquitous in nature and can be isolated from soil, water, and vegetation
(Cousin 1982). The psychrotrophic population in refrigerated raw
milk includes both Gram-positive and Gram-negative genera; they
comprise representatives of Pseudomonas, Aeromonas, Acinetobacter,
Serratia, Alcaligenes, Achromobacter, Enterobacter, Flavobacterium, Klebsiella, Bacillus, Arthrobacter, Clostridium, Lactobacillus, Listeria, Staphylococcus, Corynebacterium, Microbacterium, and Micrococcus (Cousin
1982; Champagne and others 1994; Lafarge and others 2004;
Munsch-Alatossava and Alatossava 2006). The conditions during
storage and transport in refrigerated tanks cause the raw milk microbiota to change from predominantly Gram positives to predominantly Gram negatives during bacterial growth. Gram-negative
bacteria usually account for more than 90% of the microbial population in cold raw milk that has been stored (Cousin 1982).
Currently, the predominant Gram-negative microorganisms
limiting the shelf life of ultra heat-treated (UHT) processed fluid
milk at 4 ◦ C are Pseudomonas spp., especially P. fragi, P. lundensis, and P. fluorescens-like organisms (Craven and Macauley 1992;
Ternström and others 1993; Marchand and others 2009a; De
Jonghe and others 2011). Pseudomonas spp. can grow to high
numbers and can form biofilms during refrigerated storage. Many
of them produce heat-stable extracellular lipases, proteases, and
lecithinases that contribute to milk spoilage (Shah 1994; Sorhaug
Type of bacteria
Staphylococcus Bacillus
−
+
+
−
−
+
−
−
−
−
−
−
−
−
+
+
−
−
+
+
+
+
−
−
+
+
∗
∗
∗
∗
+
−
LAB
−
−
+
+
−
+
+
−
+
+
−
−
−
∗
∗
+
Enterobacteriaceae
+
−
+
+
+
+
−
−
−
−
−
+
+
∗
∗
+
Listeria
−
−
−
−
+
+
−
∗
∗
∗
∗
∗
∗
+
+
−
and Stepaniak 1997; Marchand and others 2009b). Furthermore,
many of these enzymes remain active even following thermal processing steps that destroy their producing organisms (Garcia and
others 1989; Sorhaug and Stepaniak 1997; Marchand and others
2009b). Degradation of milk components through various enzymatic activities can reduce the shelf life of processed milk. For
example, digestion of casein by proteases can lead to bitter of flavors and the clotting and gelation of milk (Chen and others 2003;
Datta and Deeth 2003). Lipases hydrolyze tributyrin and other
milk fat glycerides to yield free fatty acids, which cause milk to
taste rancid, bitter, unclean, and soapy. Lecithinases degrade milk
fat globule membrane phospholipids and increase the susceptibility of milk fat to the action of lipases (Cousin 1982; Shah 2000).
The hydrolytic products of milk fats and proteins always decrease
the organoleptic quality of fluid milk products.
In raw milk holding equipment, 2 distinct but connected phases
are available for microbial growth: the liquid phase, in which
planktonic cells proliferate, and the solid/liquid interface (such as
milk-covered cooling tank walls) where cells can attach and form
biofilms (Wong and Cerf 1995; Somers and others 2001). Each
phase constitutes a unique habitat and cells can move from one
to the other, depending on growth stage, nutrient availability, and
flow shear forces (Stoodley and others 2002). Pseudomonas spp. and
Streptococcus spp. are among the bacteria most frequently isolated
from surfaces in the food industry (Sundheim and others 1992;
Mettler and Carpentier 1998; Flint and others 1999; Flint and
others 2000; Simões and others 2008). While streptococci form
predominantly monospecies biofilms on heat exchanger plates in
the downstream side of the sections of pasteurizers (Bouman and
others 1982; Driessen and others 1984; Flint and others 1999),
Pseudomonas spp. are more likely to produce multispecies biofilms
on the walls of milk cooling tanks or pipelines prior to heat processing. The development of a single-species biofilm may occur
due to the fact that heat-sensitive species are killed during pasteurization leaving only heat-resistant species such as Streptococcus bovis
and Streptococcus thermophilus (Bouman and others 1982; Flint and
others 2000). In addition to the risk of being a severe contamination source to subsequent milk batches passing the biofilm region,
Pseudomonas biofilms may attract and/or shelter other (spoilage or
pathogenic) bacteria. In this regard, Simoes and others (2009) illustrated that dual biofilms of P. fluorescens and B. cereus were about
5 times more metabolically active than P. fluorescens monospecies
biofilms. In terms of viability, P. fluorescens was more tolerant to
antimicrobials than B. cereus in single-species biofilms. Moreover,
138 Comprehensive Reviews in Food Science and Food Safety r Vol. 11, 2012
c 2012 Institute of Food Technologists®
Biofilm risks in dairy practice . . .
bacteria were more susceptible to antimicrobials in single-species
biofilms than in dual-species biofilms (Simoes and others 2009).
Kives and others (2005) reported on the cocultivation of Lactococcus
lactis ssp. cremoris and Pseudomonas fluorescens in refrigerated milk.
Compared to each monospecies biofilm, the dual-species biofilms
showed a more developed structure in which both species were
maintained. The benefit was most significant for L. lactis, a poor
biofilm former, which probably benefited from the enhanced attaching potential provided by the quickly developing matrix originating from P. fluorescens. In addition, the latter strain consumed
much of the available oxygen in the biofilm, which was an additional advantage for the anaerobic L. lactis. In return, P. fluorescens
utilized the lactic acid produced by L. lactis as a nutrient source.
This interdependence led to compact masses of P. fluorescens entrapping L. lactis cells. Besides, P. fragi has been shown to enhance
the attachment of L. monocytogenes to glass surfaces (Sasahara and
Zottola 1993). The enhancement was attributed to polysaccharide production by P. fragi. Also, Flavobacterium spp. have been
shown to promote biofilm formation of L. monocytogenes (Bremer and others 2001). Probably, P. fragi and Flavobacterium spp.
act as primary colonizers of the surface, making adhesion easier
for L. monocytogenes. Lindsay and others (2002) demonstrated an
enhancement of B. cereus cell attachment by 0.5–1 log cfu/cm2
in a binary biofilm with P. fluorescens. In return, B. cereus appeared to protect P. fluorescens from the sanitizers used in this
study.
Another important feature of milk-spoiling Pseudomonas
biofilms might be the altered phenotype of the inhabiting
strains. One process involved in phenotypic diversification is
phase variation, which is usually a reversible, high-frequency
phenotype switching corresponding to differential expression of
one or several genes. The genes implicated in phase variations
encode the GacA/GacS 2-component regulatory system (van den
Broek and others 2005b), which regulates secondary metabolism,
exo-enzyme production, quorum sensing, motility, and, not
surprisingly, biofilm formation (Lapouge and others 2008).
Phase variation, which is inducible by environmental factors
such as temperature (Schwan and others 1992; Gally and others
1993), medium composition (White-Ziegler and others 2000),
and stress conditions (White-Ziegler and others 2002), can
influence the growth characteristics and extracellular enzyme
production in Pseudomonas spp. (Chabeaud and others 2001; van
den Broek and others 2005a). Since phase variation seems to be
induced in the biofilm growth mode, this can hold important
implications toward the production of milk-spoiling enzymes by
pseudomonads present in dairy biofilms. Workentine and others
(2010) characterized 2 distinct colony morphology variants from
biofilms of P. fluorescens mutants missing the GacS sensor kinase.
These variants produced more biofilm cell mass and displayed
a change in amino acids and metabolites produced through
glutathione biochemistry. In laboratories, these types of colony
morphology variants are recovered at increasing frequencies
when biofilms are exposed to stressors such as oxidative agents,
antibiotics, and metal ions (Davies and others 2007; Harrison and
others 2007; Boles and Singh 2008). This suggests that phenotypic
switching might play an important role in the survival of a biofilm
population during environmental stresses. Since biofilms are frequently exposed to sanitizers during cleaning of dairy processing
equipment, phenotypic switching may occur on a regular basis
and even influence the enzyme production by pseudomonads
adding an additional spoiling factor to the subsequently processed
milk batch. This might certainly be the case if such Pseudomonas
c 2012 Institute of Food Technologists®
biofilms are present in the cooling equipment in farms or holding
silos in the dairy factory. Since the enzymes might be released
from the biofilms into the milk, without bacterial detachment,
the contamination might go unnoticed until problems arise with
the shelf life of the heat-treated dairy products. This might be of
special importance to UHT-processed milk since the Pseudomonas
enzymes are heat-resistant and withstand the heating conditions
applied (Dogan and Boor 2003; Marchand and others 2009b).
Therefore, it is clear that dairy processing equipment should be
checked regularly for biofilm formation and cleaned efficiently
in order to prevent milk-spoiling events or consumer exposure to
pathogens.
Efficacy of Different Cleaners and Sanitizers on Dairy
Biofilms
Biofilm control in dairy manufacturing plants generally involves
a process called cleaning-in-place (CIP). This cleaning process is
characterized by the cleaning of complete plant items or pipeline
circuits without the need to dismantle or open the equipment and
with little or no manual involvement from the operator (Bremer
and others 2006). CIP can be defined as circulation of cleaning
liquids through machines and other equipment in a cleaning circuit. The passage of the high-velocity flow of liquids over the
equipment surfaces generates a mechanical scouring effect that
dislodges milk deposits. This, however, only applies to the flow in
pipes, heat exchangers, pumps, valves, separators, and so on. The
normal technique for cleaning large milk storage tanks is to spray
the detergent on the upper surfaces, and then allow it to run down
the walls. The mechanical scouring is then often insufficient, but
this effect can be improved to some extent by use of specially
designed spray devices (Bylund 1995).
In the dairy industry, CIP systems generally involve the sequential use of caustic (sodium hydroxide) and acid (nitric acid)
wash steps, and chemicals originally selected for their ability to
remove organic (proteins and fat) and inorganic (calcium phosphate and other minerals) fouling layers (Kessler 1981). In some
cases, sanitizers are also incorporated in the CIP system (Kessler
1981; Bylund 1995). The choice of the cleaning process is determined by the type and composition of the soiling matter as well
as by the design of the equipment to be cleaned (Kessler 1981).
In addition, dairy CIP programs differ according to whether the
circuit to be cleaned contains heated surfaces or not. Examples of
both cleaning programs are given in Table 3. The main difference
between the 2 types is that acid circulation must always be included in the first type to remove encrusted protein and salts from
the surfaces of heat-treatment equipment. To enhance cleaning
effectiveness, caustic detergents and caustic additives have been
developed, which contain surfactants, emulsifying agents, chelating compounds, and complexing agents (Bremer and others 2006).
Traditionally, chlorine (sodium hypochlorite)-based sanitizers have
been used, however, a wide variety of sanitizers including quaternary ammonium compounds, anionic acids, iodophores, and
chlorine-based compounds are currently in use or being evaluated
for use in CIP systems (Joseph and others 2001; Parkar and others
2004; Bremer and others 2006). The selection of detergents and
disinfectants in the dairy industry depends on the efficacy, safety,
and rinsability of the agent and whether it is corrosive or affects
the sensory values of the processed products.
A feature of CIP operations, evident in both industrial and
laboratory-scale systems, is their variability in effectiveness in
eliminating surface-adherent bacteria or biofilms (Austin and
Bergeron 1995; Faille and others 2001; Dufour and others 2004).
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Biofilm risks in dairy practice . . .
This variability is not surprising as a large number of factors can
influence CIP effectiveness including the nature, age and composition of the biofilm, the cleaning agent composition and concentration, cleaning time, cleaning agent temperature, degree of
turbulence of the cleaning solution, and the characteristics of the
surface being cleaned (Stewart and Seiberling 1996; Changani and
others 1997; Lelievre and others 2001; Lelievre and others 2002).
Cleaning effectiveness is dependent on both product and processing plant-specific variables. The optimal CIP regime varies among
dairy processing plants and also over time within a given plant. In
this regard, Bremer and others (2006) have demonstrated that
dairy biofilms consisting of Gram-positive spore-forming bacilli
and thermoresistant streptococci were not adequately removed by
a standard CIP procedure (water rinse, 1.0% sodium hydroxide
at 65 ◦ C for 10 min, water rinse, 1.0% nitric acid at 65 ◦ C for
10 min, and water rinse). However, the authors found that when a
caustic additive (containing chelating and sequestering agents and
surface-active wetting agents) and a nitric acid blend (containing
surfactants) were added, a 3.8 log reduction in the number of
cells recovered from a stainless steel surface was achieved (Bremer
and others 2006). This study thus illustrated that the effectiveness
of a “standard” CIP can possibly be enhanced through testing
and use of caustic and nitric blends. Hydrogen peroxide has been
found to be effective in removing biofilms from equipment used
in hospitals (MattilaSandholm and Wirtanen 1992). Wirtanen and
others (1995) showed that the peroxide-based disinfectant was the
most effective disinfectant against Pseudomonas biofilms when the
microbiological activity was measured using conventional cultivation. The effect of hydrogen peroxide is based on the production
of free radicals, which affect the biofilm matrix. The microbicidal effect of peracetic acid on microbes in biofilms was shown
to be variable (Christensen 1989; Kramer 1997). Aldehydes did
not break the biofilm, but rather seemed to improve its stability. The biofilm must be disrupted in some way before chemical
agents such as peracetic acid and aldehydes can be used effectively (Wirtanen 2004). The effect of ozone treatments has been
found to vary depending on the processing circumstances and
the bacteria tested; ozonation proved very effective in the treatment of cooling water systems (Lin and Yeh 1993). Disinfectants
are most effective in the absence of organic (such as fat-, sugar-,
and protein-based) materials (Wirtanen 2004). Organic substances,
pH, temperature, concentration, and contact time generally control the efficacy of disinfectants (Mosteller and Bishop 1993). The
disinfectants must be effective, safe, and easy to use and also easily
rinsed off from surfaces, leaving no toxic residues or traces that
affect the sensory attributes of the food product. In a study by
Lequette and others (2010), the cleaning efficiency of polysaccharidases and proteolytic enzymes against biofilms of bacterial
species found in food industry processing lines was analyzed. Two
serine proteases and an α-amylase appeared to be the most efficient enzymes. Proteolytic enzymes promoted biofilm removal of a
larger range of bacterial species than polysaccharidases, while more
specifically, the serine proteases were more efficient in removing
Bacillus biofilms and the polysaccharidases were better at removing
P. fluorescens biofilms (Lequette and others 2010). Solubilization of
enzymes with a buffer containing surfactants and dispersing and
chelating agents enhanced the efficiency of polysaccharidases and
proteases in removing biofilms of Bacillus and P. fluorescens, respectively (Lequette and others 2010). Considering these results,
a combination of enzymes targeting several components of EPS,
surfactants, and dispersing and chelating agents could be a good
alternative to chemical cleaning agents.
Ultrasonic Cleaning
Most of the previously described cleaning and disinfection processes are well known and often used in food industry premises.
A less familiar technique applicable in cleaning off place (COP)
systems is ultrasonic cleaning. The use of ultrasound is one of the
most recently studied promising cleaning methods (Kallioinen and
Manttari 2011). Ultrasound is a form of energy generated by pressure/sound waves of frequencies that are too high to be detected
by the human ear, namely, above 16 kHz (Jayasooriya and others
2004). During a sonication process, longitudinal waves are created when a sonic wave meets a liquid medium, thereby creating
regions of alternating compression and expansion. These regions
of pressure change cause cavitation to occur, and gas bubbles are
formed in the medium. These bubbles have a larger surface area
during the expansion cycle, which increases the diffusion of gas,
causing the bubble to expand (Dolatowski and others 2007). A
point is reached where the ultrasonic energy provided is not sufficient to retain the vapor phase in the bubble; therefore, rapid
condensation occurs. The condensed molecules collide violently,
creating shock waves. Depending on the frequency used and the
sound wave amplitude applied, a number of physical, chemical,
and biochemical effects can be observed, which enable a variety
of applications. In ultrasonic cleaning, biofilm or foulant removal
takes place as a result of mechanical actions, caused by ultrasound
in the fluid medium, or as a result of chemical interactions of
foulants with radicals, which are generated into the liquid through
ultrasonic treatment (Ashokkumar and Grieser 1999; Lamminen
and others 2004; Kallioinen and Manttari 2011). In the study
by Oulahal and others (2004), 2 ultrasonic devices, a flat (T1)
and a curved (T2) ultrasonic transducers, were developed to remove biofilms from opened and closed surfaces, respectively. The
authors obtained total removal of Escherichia coli and Staphylococcus aureus milk model biofilms with the T1 transducer (10 s at
40 kHz), while the T2 transducer failed to completely remove
these model biofilms: 30% and 60% removal for the E. coli and S.
aureus biofilms, respectively (Oulahal and others 2004). When a
chelating agent was combined with the ultrasound of transducer
T2, complete removal was obtained in the E. coli biofilm, but
no enhancement could be obtained in the S. aureus milk biofilm
(Oulahal and others 2004). In the study by Baumann and others
(2009), the efficacy of ultrasound and ozonation was determined
using for the removal of L. monocytogenes biofilms from stainless
steel chips. Ultrasound (20 kHz, 100% amplitude, 120 W) was
applied for 30 or 60 s at a distance of 2.54 cm from a biofilm
chip, while it was submerged in 250 mL of sterile potassium phosphate buffer (pH 7.0). Ozone was cycled through the 250 mL of
potassium phosphate buffer containing the biofilm chip also for
30 or 60 s at concentrations of 0.25, 0.5, or 1.0 ppm. Each of the
treatments alone resulted in a significant detachment of the cells,
with ultrasound being the most effective. For the ozone in combination with ultrasound treatment, detachment was higher than
by either treatment alone (Baumann and others 2009). It can be
questioned that ultrasound is usable in large plants used for milk
powder, cheese, or yogurt because of the necessary scaling-up.
Nevertheless, for certain applications (such as cleaning of storage
tanks) in milk processing, ultrasound may be a fruitful option.
Biofilm Resistance to Antimicrobial Agents
Bacteria in biofilms have intrinsic mechanisms that protect them
from even the most aggressive environmental conditions, including
the exposure to antimicrobials (Gilbert and others 2002; Cloete
2003; Davies 2003). Dynes and others (2009) investigated the
140 Comprehensive Reviews in Food Science and Food Safety r Vol. 11, 2012
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Biofilm risks in dairy practice . . .
effect of subinhibitory concentrations of 4 antimicrobial agents.
Their results indicate that each antimicrobial agent elicited a
unique response: P. fluorescens cells and biofilms changed their morphology and architecture, as well as the distribution and abundance
of biomacromolecules, in particular the exopolymer matrix. Diversity in microbial communities leads to a variety of complex
relationships involving inter- and intraspecies interactions (Berry
and others 2006; Hansen and others 2007). The surface colonization by one type of bacterium can enhance the attachment of
others to the same surface. This process allows the development of
multispecies communities often possessing greater combined stability and resilience than that of each individual species (Moller and
others 1998; Burmolle and others 2006). In this regard, Norwood
and Gilmour (2000) investigated the effect of sodium hypochlorite on multispecies biofilms containing P. fragi, S. xylosus, and L.
monocytogenes. In a constant-depth film fermenter, in the absence
of sodium hypochlorite, the steady-state population of L. monocytogenes was only 1.5% of the total plate count, while the P. fragi
proportion amounted to 59% and was significantly greater than
that of S. xylosis (39.5%), showing a greater competitive advantage
for the pseudomonad in the unchallenged biofilm. While all 3
planktonic cultures, subjected to 10 ppm free chlorine for 30 s,
were completely eliminated, only a 2 log reduction in L. monocytogenes cells in the multispecies biofilm could be achieved after
the biofilm was exposed to 1000 ppm free chlorine for 20 min.
Their study confirmed that multispecies biofilms increased protective properties over monospecies biofilms. The authors attributed
these observations to the shielding effect of increased numbers
(or aggregation) of microorganisms but also to the production of
greater amounts of EPS. Sommer and others (1999) also found
an increase of Pseudomonas biofilm resistance to chlorine with increasing age of the biofilm. Here, it was speculated that metabolic
change or the production of exocellular compounds might be responsible for the interaction with free chlorine or prevention of its
diffusion in the biofilm. Also, Lindsay and others (2002) reported
an increased resistance of P. fluorescens against a chlorine dioxide
containing sanitizer through protection by the mere presence of
the more tolerant B. cereus. Such shielding results from the physical
protection or engulfment of the sensitive species by the tolerant
one. Modifications in EPS composition and quantity also appear
to influence bacterial resistance. Certainly, in pseudomonads, the
capacity to alter EPS composition may be part of its intrinsic
resistance to antimicrobials (Dynes and others 2009).
Monitoring, Detection, and Lab-Scale Biofilm Research
The bacterial enumeration of biofilms helps in identifying the
type(s) of microorganisms involved in biofilm formation. The
different methods employed for sampling and enumeration of
biofilms in a dairy plant are swabbing, rinsing, agar flooding, and
agar contact methods (Kumar and Anand 1998). The organisms
found in biofilms, however, are not always easily cultured, resulting
in an underestimate—or no detection at all—of the true biofilm
population inside a liquid handling system (Wirtanen 1995). As an
alternative, scraping (Frank and Koffi 1990), vortexing (Mustapha
and Liewen 1989), and ultrasound are often used. Studies have
shown that ultrasound generates sufficient cavitational bubble activity to remove biofilms from metal, glass, ceramic, and plastic
surfaces (Stickler and Hewett 1991). The ultrasonic treatment for
microbial recovery consists of immersing and agitating samples,
usually in glass test tubes or flasks containing liquid, in a highfrequency ultrasonic bath (18–55 kHz) (Jeng and others 1990).
The disadvantage of this lab-scale method, however, is that it is
c 2012 Institute of Food Technologists®
an invasive technique. Surface samples have to be cut out from
the plant and studied in bath sonicators (Oulahal-Lagsir and others 2000). Besides, grooves, crevices, dead, ends, and corrosion
patches are areas where biofilms readily occur but are hard to access, thus hampering sampling of such areas. In addition, some
of the bacteria in biofilms in dairy environments are subjected to
various stresses such as starvation, chemicals, heat, cold, and desiccation that may injure the cells and render them unculturable. Fortunately, a combination of disciplines can be used to develop data
on biofouling in liquid operations. The biofouling monitoring systems can provide different levels of information according to their
specific design (Flemming 2003; Tamachkiarow and Flemming
2003). For example, some are able to assess the biofilm dynamics,
attachment/detachment events, but cannot differentiate between
the constituents of such layers (such as biotic/abiotic) (Pereira and
Melo 2009). More specific monitoring devices are able to characterize the chemical/biological composition of a given fouling
layer, although they are too sophisticated and costly to be operated in an industrial setting (Pereira and Melo 2009). For example,
to monitor the onset of buildup of fouling on internal surfaces of
heat exchangers, a commercial sensor can be used to measure the
local heat flux and temperature on the hot side of a plate-type heat
exchanger. A real-time estimate of the fouling rate can be obtained
by calculating the heat transfer coefficient normalized to its value
at the beginning of the run (Bennet 2007). Another system to evaluate global fouling can be used by calculating the energy balance
over a tubular-type heat exchanger. In that case, the overall heat
transfer coefficient is calculated by measuring the inlet and outlet temperatures (Bennet 2007). Also, specular reflectance Fourier
transform infrared (FTIR) spectroscopy can capture a fingerprint
of the organic constituents of a fouling film and epifluorescence
optical microscopy with the appropriate fluorescent nucleic acid
stains can be used to determine whether the pipe wall fouling
is biological or abiotic in nature. Different methods are available
that report biofilm growth online, in real-time and nondestructively, but they all are based on physical methods. One example is a
method that uses 2 turbidity measurement devices, one of which is
constantly cleaned. The difference of signals is proportional to the
biomass developing on the noncleaned window (Klahre and Flemming 2000). Another one is the fiber active device (FOS), which is
based on a light fiber integrated in the test surface, measuring the
scattered light of material deposited on the tip (Tamachkiarow and
Flemming 2003). Fornalik (2008), on the other hand, has developed a fouling cell assembly in 316L-grade stainless steel that may
be placed in dairy pipes and silos. Such assemblies enable monitoring of biofilm development without removal of the processing
equipment out of the plant and can be used to generate objective
data on the effectiveness of cleaning procedures (Fornalik 2008).
Due to the limitations of studying biofilm development in practical settings, most of biofilm research has been performed in
laboratory-based model systems. Despite the many different types
of biofilm model systems described in the scientific literature, none
of them can be considered as the optimal, universally applicable
model system. On the contrary, every researcher has to choose a
particular model system that enables to address the specific research
questions formulated in the beginning of the study. Since biofilm
growth simulating devices are beyond the scope of this review,
only a small oversight is given here (Table 2). For further information on this topic, the reader is directed toward other extensive
reviews (McLandsborough and others 2006).
After selection of an appropriate growth model to produce
biofilms, the need arises to reliably quantify the number of cells in
Vol. 11, 2012 r Comprehensive Reviews in Food Science and Food Safety 141
Biofilm risks in dairy practice . . .
Table 2–Biofilm growth simulating devices. Primary use and limitations in biofilm research, adapted from McLandsborough and others (2006).
Device
Microtiter plate∗
BioFlux
Polycarbonate
membranes∗
Capillary reactor
Flow cell reactor∗
Robbin’s device∗
Modified Robbin’s device
Calgary biofilm device
Rotating disk reactor
CDC biofilm reactor
Rotating annular reactor∗
Batch and batch-fed
growth system∗
Constant-depth film
fermentor
Animal models
Primary Use
Limitations
Useful in genetic studies because of
high throughput screening of
“static biofilms.”
Useful in genetic studies because of
high troughput screening of “flow
biofilms.”
Simple methodology and easy to use.
Suitable for antimicrobial
penetration tests.
Biofilm structure is formed on glass
capillary. Direct microscopic
observation is feasible.
Biofilms can be studied under either
laminar or turbulent flow in order
to simulate the changes in fluid
velocity that occur during the
operation of industrial reactors.
Using a brass pipe, removable
sections of the wall can be
removed to test biofilm growth.
Used in industrial biofouling.
Allows several materials to be tested.
Used in industrial biofouling.
High troughput. Rapid and
reproducible assays in biofilm
susceptibility to antibiotics.
Different biomaterials can be used for
colonization and shear forces can
be controlled.
Used to follow biofilm formation
(under moderate to high shear),
characterize biofilm structure, and
assess the effect of antimicrobial
agents.
Application of a well defined shear
field
Suitable for a wide variety of biofilm
experiments
Only for early stages of biofilm
formation.
Biofilm growth and resistance to
antimicrobials in multispecies
biofilms
Biofilm formation and distribution in
tissues can be monitored. Used in a
biofilm model of chronic cystitis
and prostatitis.
In order to reach a steady-state
biofilm, the biofilm has to be grown
in a chemostat.
Time-consuming and regulatory
issues
References
Limited to GFP-expressing bacteria.
(O’Toole and Kolter 1998; Djordjevic
and others 2002; Stepanovic and
others 2004)
(Benoit and others 2010)
Since bacterial cultures are manually
deposited on the membrane,
biofilms do not naturally develop.
Biofilm growth is limited to a single
surface.
(Anderl and others 2000; Werner and
others 2004; Borriello and others
2004; Tang 2011)
(Mccoy and others 1981; Werner and
others 2004)
None for the purpose the method was
designed for.
(Pereira and others 2002; Simões and
others 2008)
Just one type of material can be
tested at a time.
(Mccoy and others 1981; Mittelman
1998)
Used for traditional biofilm cultures
and not for genetic investigations.
None for the purpose the method was
designed for.
(Nickel and others 1985)
High variability seen in biofilm
formation between samples.
(Okabe and others 1999)
The baffle rotation speed has to be
carefully controlled.
(Donlan and others 2004; Goeres and
others 2005)
None for the purpose the method was
designed for
High variability seen in biofilm
formation between samples
(Camper and others 1996; Jang and
others 2006)
(Cerca and others 2004;
Sirianuntapiboon and others
2005)
(Knowles and others 2005)
(Ceri and others 1999)
(Kadurugamuwa and others 2004)
∗ Already used in milk biofilm studies.
Table 3–Examples of dairy CIP programs, adapted from Bylund (1995)
CIP wash steps for circuits with pasteurizers and other equipment with
heated surfaces (UHT, and others)
1
Rinsing with warm water for about 10 min.
2
Circulation of an alkaline detergent solution (0.5%–1.5%) for about
30 min at 75 ◦ C.
3
Rinsing out alkaline detergent with warm water for about 5 min.
4
Circulation of (nitric) acid solution (0.5%–1.0%) for about 20 min at
70 ◦ C.
5
Postrinsing with cold water.
6
Gradual cooling with cold water for about 8 min.
CIP wash steps for circuits with pipe systems, tanks, and other process
equipment with no heated surfaces
1
Rinsing with warm water for 3 min.
2
Circulation of a 0.5%–1.5% alkaline detergent at 75 ◦ C for about
10 min.
3
Rinsing with warm water for about 3 min.
4
Disinfection with hot water 90–95 ◦ C for 5 min.
5
Gradual cooling with cold tap water for about 10 min (normally no
cooling for tanks).
the developed biofilm and to determine the structure and composition of the biofilm. Biofilm structure development has been
analyzed using light, fluorescence, differential interference contrast
(DIC), transmission electron (TE), scanning electron (SE), atomic
force (AF), and confocal laser scanning microscopy (CLSM) (Ceri
and others 1999; Storgards and others 1999a; Djordjevic and others 2002; Donlan and Costerton 2002; Hunter and Beveridge
2005; Lagace and others 2006; Sigua and others 2010; Shaheen
and others 2010). CLSM has been developed in the 1980s and
allows examination of biofilms without the limitations imposed
by scanning electron microscopy (SEM) and transmission electron microscopy (TEM). Fully hydrated biofilms are analyzed by
progressive laser scans at different focal planes within the sample. Computer analysis of the scanned images permits recreation
of the 3-dimensional structure of the biofilm. The application of
CLSM combined with a number of fluorescent stains provides
an important and effective tool to analyze the composition and
structure of hydrated biofilms in situ, nondestructively, and in real
time (Lawrence and Neu 1999; Manz and others 1999). Viability
and distribution of cells within the biofilm may be analyzed as
well. When using epifluorescence or CLSM, the choice of suitable fluorescent stains is critical in order to increase the contrast
between organisms and the exopolymers in the biofilms. Nucleic
acid stains such as 4,6-diamino-2-phenylindole (DAPI) or acridin
orange have been used to stain the DNA of cells regardless of
their viability (Trachoo 2003). Other dyes sensitive to viable cells
such as propidium iodine (PI) or 5-cyano-2,3-ditolyl tetrazolium
142 Comprehensive Reviews in Food Science and Food Safety r Vol. 11, 2012
c 2012 Institute of Food Technologists®
Biofilm risks in dairy practice . . .
chloride may be used to further resolve viable and dead cells (Don- Anonymous. 2007. Ministrieel besluit houdende goedkeuring van het
document opgesteld door de erkende inteprofessionele organismen
lan and Costerton 2002).
Conclusions and Research Perspectives
Biofilms are one of the main recontamination sources of milk. It
has been established that for each planktonic bacterium detected,
there might be close to 1000 organisms present in biofilms. In
the dairy industry, mono- as well as multispecies biofilms can occur. Pathogenic bacteria can coexist within a biofilm with other
environmental organisms; an example of this is L. monocytogenes
surviving in Pseudomonas biofilms. Biofilms are difficult to remove
from milk processing environments due to the production of EPS
materials and the difficulties associated with cleaning complex
processing equipment and processing environments. Since stringent cleaning protocols are available, cleaning procedures should
be accurately applied, and ideally, the cleaning efficiency should be
evaluated. There is far too little knowledge on persisting contamination sources and existing innovative cleaning and disinfection
techniques. Therefore, it is important that research results in this
area are thoroughly communicated with the industry. In addition,
an objective “cleaning efficiency measuring system” should be developed, which in the end can lead to the issuance of directives for
economical and technical optimalization of existing CIP systems.
Biofilm control relies in the end on the design of storage and processing equipment, effective cleaning and sanitizing procedures,
and the correct implementation and application. The management of these factors is important to ensure safe and good-quality
milk and dairy products.
From a practical viewpoint, future research could also focus
on coating strategies to reduce microbial attachment on dairy
equipment and cleaners and sanitizers with fortified properties
(for example, addition of EPS or protein-degrading enzymes). In
addition, exploration of the exo-enzyme production by biofilm
pseudomonads might be of interest when milk spoilage is under study. From a more fundamental research approach, it could
be very useful to investigate the temperature effect on the development of Pseudomonas biofilms. Since both biofilm production and exo-enzyme production are under the same control in
pseudomonads and exo-enzyme production is elevated at lower
temperatures (certainly in milk), it could be interesting to check
if milk Pseudomonas strains have a competitive advantage over the
other milk bacteria due to an increased biofilm-forming capacity
at lower temperatures.
Acknowledgments
The authors wish to thank the Agency for Innovation by Science
and Technology (IWT) for financial support.
References
Agarwal S, Sharma K, Swanson BG, Yuksel GU, Clark S. 2006. Nonstarter
lactic acid bacteria biofilms and calcium lactate crystals in cheddar cheese.
J Dairy Sci 89(5):1452–66.
Anderl JN, Franklin MJ, Stewart PS. 2000. Role of antibiotic penetration
limitation in Klebsiella pneumoniae biofilm resistance to ampicillin and
ciprofloxacin. Antimicrob Agents Chemother 44(7):1818–24.
Anonymous. 2004. Regulations on materials intended to come into contact
with food EC1935/2004. Off J Eur Union L338:1–17.
Anonymous. 2005. Microbiological criteria for food products EC2073/2005.
Off J Eur Union L338:1–74.
Anonymous. 2006. Hygiene directives for foods of animal origin EC1662/
2006. Off J Eur Union L320:1–10.
c 2012 Institute of Food Technologists®
betreffende de modaliteiten van de controle van de kwaliteit van de rauwe
koemelk. Belgisch Staatsblad 824:7679–83.
Anonymous. 2011. DQA—dairy quality assurance scheme regulations for
Belgium. (Lastenboek IKM Productie) Version 5-11-86.
Ashokkumar M, Grieser F. 1999. Ultrasound assisted chemical processes.
Rev Chem Eng 15(1):41–83.
Austin JW, Bergeron G. 1995. Development of bacterial biofilms in dairy
processing lines. J Dairy Res 62(3):509–19.
Baier RE. 2006. Surface behaviour of biomaterials: the theta surface for
biocompatibility. J Mater Sci—Mater Med 17(11):1057–62.
Baumann AR, Martin SE, Hao F. 2009. Removal of Listeria monocytogenes
biofilms from stainless steel by use of ultrasound and ozone. J Food Prot
72(6):1306–9.
Bayles KW. 2007. The biological role of death and lysis in biofilm
development. Nat Rev Microbiol 5(9):721–6.
Bennet H. 2007. Aspects of fouling in dairy processing: a thesis presented in
partial fulfillment for the degree of Doctor of Philosophy in Food
Engineering at Massey Univ., Palmerston North, New-Zealand; 1–172.
Benoit MR, Conant CG, Ionescu-Zanetti C, Schwartz M, Matin A. 2010.
New device for high-throughput viability screening of flow biofilms. Appl
Environ Microbiol 76(13):4136–42.
Bernbom N, Ng YY, Jorgensen RL, Arpanaei A, Meyer RL, Kingshott P,
Vejborg RM, Klemm P, Gram L. 2009. Adhesion of food-borne bacteria to
stainless steel is reduced by food conditioning films. J Appl Microbiol
106(4):1268–79.
Berry D, Xi CW, Raskin L. 2006. Microbial ecology of drinking water
distribution systems. Curr Opin Biotechnol 17(3):297–302.
Bjarnsholt T, Jensen PO, Fiandaca MJ, Pedersen J, Hansen CR, Andersen
CB, Pressler T, Givskov M, Hoiby N. 2009. Pseudomonas aeruginosa biofilms
in the respiratory tract of cystic fibrosis patients. Pediatr Pulmonol
44(6):547–58.
Blackman IC, Frank JF. 1996. Growth of Listeria monocytogenes as a biofilm
on various food-processing surfaces. J Food Prot 59(8):827–31.
Boles BR, Singh PK. 2008. Endogenous oxidative stress produces diversity
and adaptability in biofilm communities. Proc Natl Acad Sci USA
105(34):12503–8.
Borriello G, Werner E, Roe F, Kim AM, Ehrlich GD, Stewart PS. 2004.
Oxygen limitation contributes to antibiotic tolerance of Pseudomonas
aeruginosa in biofilms. Antimicrob Agents Chemother 48(7):2659–64.
Boulangé-Petermann L, Rault J, Bellon-Fontaine MN. 1997. Adhesion of
Streptococcus thermophilus to stainless steel with different surface topography
and roughness. Biofouling 11:201–16
Bouman S, Lund DB, Driessen FM, Schmidt DG. 1982. Growth of
thermoresistant Streptococci and deposition of milk constituents on plates of
heat-exchangers during long operating times. J Food Prot 45(9):806–12.
Bremer PJ, Fillery S, McQuillan AJ. 2006. Laboratory-scale clean-in-place
(CIP) studies on the effectiveness of different caustic and acid wash steps on
the removal of dairy biofilms. Intl J Food Microbiol 106(3):254–62.
Bremer PJ, Monk I, Osborne CM. 2001. Survival of Listeria monocytogenes
attached to stainless steel surfaces in the presence or absence of Flavobacterium
spp. J Food Prot 64(9):1369–76.
Bremer PJ, Seale B, Flint S, Palmer J. 2009. Biofilms in dairy processing. In:
Fratamico PM, Annous BA, Gunther NW, IV, editors. Biofilms in the food
and beverage industries. Oxford, Cambridge, New Delhi: Wood head
Publishing Limited. p 396–431.
Brooks JD, Flint SH. 2008. Biofilms in the food industry: problems and
potential solutions. Intl J Food Sci Technol 43(12):2163–76.
Burgess SA, Lindsay D, Flint SH. 2010. Thermophilic bacilli and their
importance in dairy processing. Intl J Food Microbiol 144(2):215–25.
Burmolle M, Webb JS, Rao D, Hansen LH, Sorensen SJ, Kjelleberg S. 2006.
Enhanced biofilm formation and increased resistance to antimicrobial agents
and bacterial invasion are caused by synergistic interactions in multispecies
biofilm. Appl Environ Microbiol 72(6):3916–23.
Bylund G. 1995. Cleaning of dairy equipment. In: Dairy processing
handbook. Lund, Swedeb: Tetra Pak Processing Systems. p 403–14.
Camper AK, Jones WL, Hayes JT. 1996. Effect of growth conditions and
substratum composition on the persistence of coliforms in
mixed-population biofilms. Appl Environ Microbiol 62(11):4014–8.
Cappello S, Guglielmino SPP. 2006. Effects of growth temperature on
polystyrene adhesion of Pseudomonas aeruginosa ATCC 27853. Brazilian J
Microbiol 37(3):205–7.
Vol. 11, 2012 r Comprehensive Reviews in Food Science and Food Safety 143
Biofilm risks in dairy practice . . .
Cerca N, Pier GB, Vilanova M, Oliveira R, Azeredo J. 2004. Influence of
batch or fed-batch growth on Staphylococcus epidermidis biofilm
formation. Lett Appl Microbiol 39(5):420–4.
Ceri H, Olson ME, Stremick C, Read RR, Morck D, Buret A. 1999. The
calgary biofilm device: new technology for rapid determination of antibiotic
susceptibilities of bacterial biofilms. J Clin Microbiol 37(6):1771–6.
Chabeaud P, De Groot A, Bitter W, Tommassen J, Heulin T, Achouack W.
2001. Phase variable expression of an operon encoding extracellular alkaline
protease, a serine protease homolog, and lipase in Pseudomonas brassicacearum.
J Bacteriol 183:2117–20.
Chambless JD, Stewart PS. 2007. A three-dimensional computer model
analysis of three hypothetical biofilm detachment mechanisms. Biotechnol
Bioeng 97(6):1573–84.
Champagne CP, Laing RR, Roy D, Mafu AA. 1994. Psychrotrophs in dairy
products: their effect and their control. Crit Rev Food Sci Nutr 34:1–30.
Chandy JP, Angles ML. 2001. Determination of nutrients limiting biofilm
formation and the subsequent impact on disinfectant decay. Water Res
35(11):2677–82.
Changani SD, BelmarBeiny MT, Fryer PJ. 1997. Engineering and chemical
factors associated with fouling and cleaning in milk processing. Exp
Thermal Fluid Sci 14(4):392–406.
Chen L, Daniel RM, Coolbear T. 2003. Detection and impact of
protease and lipase activities in milk and milkpowders. Intl Dairy J 7(8–9):
255–75.
Christensen BE. 1989. The role of extracellular polysaccharides in biofilms.
J Biotechnol 10(3–4):181–201.
Cloete TE. 2003. Resistance mechanisms of bacteria to antimicrobial
compounds. Intl Biodeter Biodegrad 51(4):277–82.
Costerton JW, Lewandowski Z, Caldwell DE, Korber DR, Lappinscott HM.
1995. Microbial biofilms. Annu Rev Microbiol 49:711–45.
Costerton W, Veeh R, Shirtliff M, Pasmore M, Post C, Ehrlich G. 2003.
The application of biofilm science to the study and control of chronic
bacterial infections. J Clin Invest 112(10):1466–77.
Cousin MA. 1982. Presence and activity of psychrotrophic microorganisms
in milk and dairy products: a review. J Food Prot 45:172–207.
Craven HM, Macauley BJ. 1992. Microorganisms in pasteurized milk after
refrigerated storage 1. Identification of types. Aust J Dairy Technol
47(5):38–45.
Czechowski MH. 1990. Bacterial attachment to buna-N gaskets in milk
processing equipment (reprinted). Aust J Dairy Technol 45(2):113–4.
Datta N, Deeth HC. 2003. Diagnosing the cause in proteolysis in UHT milk.
Lebensmittel-Wissenschaft und-Technol—Food Sci Technol 36:173–82.
Davies D. 2003. Understanding biofilm resistance to antibacterial agents. Nat
Rev Drug Discov 2(2):114–22.
Davies JA, Harrison JJ, Marques LLR, Foglia GR, Stremick CA, Storey DG,
Turner RJ, Olson ME, Ceri H. 2007. The GacS sensor kinase controls
phenotypic reversion of small colony variants isolated from biofilms of
Pseudomonas aeruginosa PA14. Fems Microbiol Ecol 59(1):32–46.
De Jong P. 1997. Impact and control of fouling in milk processing. Trends
Food Sci Technol 8(12):401–5.
De Jonghe V, Coorevits A, Van Hoorde K, Messens W, Van Landschoot A,
De Vos P, Heyndrickx M. 2011. Influence of storage conditions on the
growth of Pseudomonas species in refrigerated raw milk. Appl Environ
Microbiol 77(2):460–70.
Di Bonaventura G, Piccolomini R, Paludi D, D’Orio V, Vergara A, Conter
M, Ianieri A. 2008. Influence of temperature on biofilm formation by
Listeria monocytogenes on various food-contact surfaces: relationship with
motility and cell surface hydrophobicity. J Appl Microbiol 104(6):1552–61.
Dickson JS, Koohmaraie M. 1989. Cell-surface charge characteristics and
their relationship to bacterial attachment to meat surfaces. Appl Environ
Microbiol 55(4):832–6.
Djordjevic D, Wiedmann M, McLandsborough LA. 2002. Microtiter plate
assay for assessment of Listeria monocytogenes biofilm formation. Appl Environ
Microbiol 68(6):2950–8.
Dogan B, Boor KJ. 2003. Genetic diversity and spoilage potentials among
Pseudomonas spp. isolated from fluid milk products and dairy processing
plants. Appl Environ Microbiol 69(1):130–8.
Dolatowski Z, Stadnik J, Stasiak D. 2007. Application of ultrasound in food
technology. Acta Sci Polonorum, Technol Aliment 6(3):89–99.
Donlan RM, Costerton JW. 2002. Biofilms: survival mechanisms of clinically
relevant microorganisms. Clin Microbiol Rev 15(2):167–93.
Donlan RM, Piede JA, Heyes CD, Sanii L, Murga R, Edmonds P, El-Sayed
I, El-Sayed MA. 2004. Model system for growing and quantifying
Streptococcus pneumoniae biofilms in situ and in real time. Appl Environ
Microbiol 70(8):4980–8.
Driessen FM, Devries J, Kingma F. 1984. Adhesion and growth of
thermoresistant Streptococci on stainless-steel during heat-treatment of
milk. J Food Prot 47(11):848–52.
Dufour M, Simmonds RS, Bremer PJ. 2004. Development of a laboratory
scale clean-in-place system to test the effectiveness of “natural”
antimicrobials against dairy biofilms. J Food Prot 67(7):1438–43.
Dynes JJ, Lawrence JR, Korber DR, Swerhone GDW, Leppard GG,
Hitchcock AP. 2009. Morphological and biochemical changes in
Pseudomonas fluorescens biofilms induced by sub-inhibitory exposure to
antimicrobial agents. Can J Microbiol 55(2):163–78.
EFSA. 2008. Guidance document on the submission of a dossier on a
substance to be used in food contact materials for evaluation by EFSA by the
pannel on additives, flavourings, processing aids and materials in in contact
with food (AFC). Eur Food Saf Authority 1–125.
Eneroth A, Christiansson A, Brendehaug J, Molin G. 1998. Critical
contamination sites in the production line of pasteurised milk, with
reference to the psychrotrophic spoilage flora. Intl Dairy J 8(9):829–34.
Faille C, Carpentier B. 2009. Food contact surfaces, surface soiling and
biofilm formation. In: Fratamico PM, Annous BA, Gunther NW, IV,
editors. Biofilms in the food and beverage industries. Oxford, Cambridge,
New Delhi: Wood head Publishing Limited. p 304–30.
Faille C, Fontaine F, Benezech T. 2001. Potential occurrence of adhering
living Bacillus spores in milk product processing lines. J Appl Microbiol
90(6):892–900.
FDA. 2007. Determining the regulatory status of components of a food
contact material. U.S. Food and Drug Administration.
Flemming HC. 2003. Role and levels of real-time monitoring for successful
anti-fouling strategies—an overview. Water Sci Technol 47(5):1–8.
Flint S, Palmer J, Bloemen K, Brooks J, Crawford R. 2001. The growth of
Bacillus stearothermophilus on stainless steel. J Appl Microbiol 90(2):151–7.
Flint SH, Bremer PJ, Brooks JD. 1997a. Biofilms in dairy manufacturing
plant—description, current concerns and methods of control. Biofouling
11(1):81–97.
Flint SH, Brooks JD, Bremer PJ. 1997b. The influence of cell surface
properties of thermophilic streptococci on attachment to stainless steel.
J Appl Microbiol 83(4):508–17.
Flint SH, Brooks JD, Bremer PJ. 2000. Properties of the stainless steel
substrate, influencing the adhesion of thermo-resistant streptococci. J Food
Eng 43(4):235–42.
Flint SH, van den Elzen H, Brooks JD, Bremer PJ. 1999. Removal and
inactivation of thermo-resistant streptococci colonising stainless steel. Intl
Dairy J 9(7):429–36.
Florjanic M, Kristl J. 2011. The control of biofilm formation by
hydrodynamics of purified water in industrial distribution system. Intl J
Pharm 405(1–2):16–22.
Fornalik M. 2008. Detecting biofouling in food processing sytems.
Phototonics Spectra 58:60–1.
Frank JF, Koffi RA. 1990. Surface-adherent growth of Listeria monocytogenes
is associated with increased resistance to surfactant sanitizers and heat. J Food
Prot 53(7):550–4.
Gally DL, Bogan JA, Eisenstein BI, Blomfeld IC. 1993. Environmental
regulation of the fim switch controlling type 1 fimbrial phase variation in
Escherichia coli K-12: effects of temperature and media. J Bacteriol
175:6186–93.
Garcia ML, Sanz B, Garciacollia P, Ordonez JA. 1989. Activity and
thermostability of the extracellular lipases and proteinases from
pseudomonads isolated from raw milk. Milchwissenschaft—Milk Sci Intl
44(9):547–9.
Garny K, Horn H, Neu TR. 2008. Interaction between biofilm
development, structure and detachment in rotating annular reactors.
Bioprocess Biosyst Eng 31(6):619–29.
Gibson H, Taylor JH, Hall KE, Holah JT. 1999. Effectiveness of cleaning
techniques used in the food industry in terms of the removal of bacterial
biofilms. J Appl Microbiol 87(1):41–8.
Giffel MCT, Beumer RR, Langeveld LPM, Rombouts M. 1997. The role of
heat exchangers in the contamination of milk with Bacillus cereus in dairy
processing plants. Intl J Dairy Technol 50(2):43–7.
Gilbert P, Allison DG, Mcbain AJ. 2002. Biofilms in vitro and in vivo: do
singular mechanisms imply cross-resistance? J Appl Microbiol 92:98S–
110S.
144 Comprehensive Reviews in Food Science and Food Safety r Vol. 11, 2012
c 2012 Institute of Food Technologists®
Biofilm risks in dairy practice . . .
Goeres DM, Loetterle LR, Hamilton MA, Murga R, Kirby DW, Donlan
RM. 2005. Statistical assessment of a laboratory method for growing
biofilms. Microbiology 151:757–62.
Guillemot G, Vaca-Medina G, Martin-Yken H, Vernhet A, Schmitz P,
Mercier-Bonin M. 2006. Shear-flow induced detachment of Saccharomyces
cerevisiae from stainless steel: influence of yeast and solid surface properties.
Colloids Surf B—Biointerfaces 49(2):126–35.
Gunduz GT, Tuncel G. 2006. Biofilm formation in an ice cream plant.
Antonie Van Leeuwenhoek Intl J Gen Mol Microbiol 89(3–4):329–36.
Hansen SK, Rainey PB, Haagensen JAJ, Molin S. 2007. Evolution of species
interactions in a biofilm community. Nature 445(7127):533–6.
Harrison JJ, Ceri H, Turner RJ. 2007. Multimetal resistance and tolerance in
microbial biofilms. Nat Rev Microbiol 5(12):928–38.
Helke DM, Somers EB, Wong ACL. 1993. Attachment of Listeria
monocytogenes and Salmonella typhimurium to stainless steel and Buna-N in the
presence of milk and individual milk components. J Food Prot 56(6):479–84.
Herald PJ, Zottola EA. 1988a. Scanning electron-microscopic examination of
Yersinia-Enterocolitica attached to stainless-steel at selected temperatures
and Ph values. J Food Prot 51(6):445–8.
Herald PJ, Zottola EA. 1988b. The use of transmission electron-microscopy
to study the composition of Pseudomonas fragi attachment material. Food
Microstruct 7(1):53–7.
Heyndrickx M, Marchand S, De Jonghe V, Smet K, Coudijzer K, De Block
J. 2010. Understanding and preventing consumer milk microbial spoilage
and chemical deterioration. In: Griffiths MW, editor. Improving the safety
and quality of milk. Oxford, Cambridge, New Delhi: Woodhead Publishing
Limited. p 97–123.
Hinton AR, Trinh KT, Brooks JD, Manderson GJ. 2002. Thermophile
survival in milk fouling and on stainless steel during cleaning. Food
Bioproducts Process 80(C4):299–304.
Hood SK, Zottola EA. 1995. Biofilms in food processing. Food Control
6(1):9–18.
Hunter RC, Beveridge TJ. 2005. Application of a pH-sensitive fluoroprobe
(C-SNARF-4) for pH microenvironment analysis in Pseudomonas aeruginosa
biofilms. Appl Environ Microbiol 71(5):2501–10.
Jang A, Szabo J, Hosni AA, Coughlin M, Bishop PL. 2006. Measurement of
chlorine dioxide penetration in dairy process pipe biofilms during
disinfection. Appl Microbiol Biotechnol 72(2):368–76.
Jayasooriya SD, Bhandari BR, Torley P, D’Arcy BR. 2004. Effect of
high-power ultrasound waves on properties of meat: a review. Intl J Food
Prop 7(2):301–19.
Jeng DK, Lin LI, Hervey LV. 1990. Importance of ultrasonication conditions
in recovery of microbial contamination from material surfaces. J Appl
Bacteriol 68(5):479–84.
Joseph B, Otta SK, Karunasagar I, Karunasagar I. 2001. Biofilm formation by
Salmonella spp. on food contact surfaces and their sensitivity to sanitizers. Intl
J Food Microbiol 64(3):367–72.
Kadurugamuwa JL, Sin LV, Yu J, Francis KP, Purchio TF, Contag PR. 2004.
Noninvasive optical imaging method to evaluate postantibiotic effects on
biofilm infection in vivo. Antimicrob Agents Chemother 48(6):2283–7.
Kallioinen M, Manttari M. 2011. Influence of ultrasonic treatment on
various membrane materials: a review. Sep Sci Technol 46(9):1388–95.
Kaplan JB, Meyenhofer MF, Fine DH. 2003. Biofilm growth and detachment
of Actinobacillus actinomycetemcomitans. J Bacteriol 185(4):1399–404.
Kaplan JB, Ragunath C, Velliyagounder K, Fine DH, Ramasubbu N. 2004.
Enzymatic detachment of Staphylococcus epidermidis biofilms. Antimicrob
Agents Chemother 48(7):2633–6.
Kessler HG. 1981. Cleaning—sanitizing—sterilizing. In: Food enfineering
and dairy technology. Freising, Germany: Publishinh House Verlag
A. Kessler. p 530–76.
Kirtley SA, Mcguire J. 1989. On differences in surface constitution of dairy
product contact materials. J Dairy Sci 72(7):1748–53.
Kives J, Guadarrama D, Orgaz B, Rivera-Sen A, Vazquez J, SanJose C. 2005.
Interactions in biofilms of Lactococcus lactis ssp cremoris and Pseudomonas
fluorescens cultured in cold UHT milk. J Dairy Sci 88(12):4165–71.
Klahre J, Flemming HC. 2000. Monitoring of biofouling in papermill
process waters. Water Res 34(14):3657–65.
Knowles JR, Roller S, Murray DB, Naidu AS. 2005. Antimicrobial action of
carvacrol at different stages of dual-species biofilm development by
Staphylococcus aureus and Salmonella enterica serovar typhimurium. Appl Environ
Microbiol 71(2):797–803.
Kokare CR, Chakraborty S, Khopade AN, Mahadik KR. 2009. Biofilm:
importance and applications. Indian J Biotechnol 8(2):159–68.
c 2012 Institute of Food Technologists®
Kramer JF. 1997. Peracetic acid: a new biocide for industrial water
applications. Mater Perform 36(8):42–50.
Krysinski EP, Brown LJ, Marchisello TJ. 1992. Effect of cleaners and
sanitizers on Listeria monocytogenes attached to product contact surfaces.
J Food Prot 55(4):246–51.
Kumar CG, Anand SK. 1998. Significance of microbial biofilms in food
industry: a review. Intl J Food Microbiol 42(1–2):9–27.
Lafarge V, Ogier JC, Girard V, Maladen V, Leveau JY, Gruss A,
Delacroix-Buchet A. 2004. Raw cow milk bacterial population shifts
attributable to refrigeration. Appl Environ Microbiol 70(9):5644–50
Lagace L, Jacques M, Mafu AA, Roy D. 2006. Compositions of maple sap
microflora and collection system biofilms evaluated by scanning electron
microscopy and denaturing gradient gel electrophoresis. Intl J Food
Microbiol 109(1–2):9–18.
Lamminen MO, Walker HW, Weavers LK. 2004. Mechanisms and factors
influencing the ultrasonic cleaning of particle-fouled ceramic membranes.
J Membr Sci 237(1–2):213–23.
Langeveld LPM, van Montfort Quasig RMGE, Weerkamp AH, Waalewijn
R, Wever JS. 1995. Adherence, growth and release of bacteria in a tube heat
exchanger for milk. Netherlands Milk Dairy J 49(4):207–20.
Lapouge K, Schubert M, Allain FHT, Haas D. 2008. Gac/Rsm signal
transduction pathway of gamma-proteobacteria: from RNA recognition to
regulation of social behaviour. Mol Microbiol 67(2):241–53.
Lawrence JR, Neu TR. 1999. Confocal laser scanning microscopy for
analysis of microbial biofilms. Biofilms 310:131–44.
Le-Clech P, Chen V, Fane TAG. 2006. Fouling in membrane bioreactors
used in wastewater treatment. J Membr Sci 284(1–2):17–53.
Lee JH, Kaplan JB, Lee WY. 2008. Microfluidic devices for studying growth
and detachment of Staphylococcus epidermidis biofilms. Biomed Microdevices
10(4):489–98.
Lelievre C, Antonini G, Faille C, Benezech T. 2002. Cleaning-in-place—
modelling of cleaning kinetics of pipes soiled by Bacillus spores assuming a
process combining removal and deposition. Food Bioproducts Process
80(C4):305–11.
Lelievre C, Faille C, Benezech T. 2001. Removal kinetics of Bacillus cereus
spores from stainless steel pipes under CIP procedure: influence of soiling
and cleaning conditions. J Food Process Eng 24(6):359–79.
Lequette Y, Boels G, Clarisse M, Faille C. 2010. Using enzymes to remove
biofilms of bacterial isolates sampled in the food-industry. Biofouling
26(4):421–31.
Liao BQ, Bagley DM, Kraemer HE, Leppard GG, Liss SN. 2004. A review
of biofouling and its control in membrane separation bioreactors. Water
Environ Res 76(5):425–36.
Lin SH, Yeh KL. 1993. Looking to treat waste water. Try ozone. Chem Eng
100(5):112–116.
Lindsay D, Brozel VS, Mostert JF, von Holy A. 2002. Differential efficacy of
a chlorine dioxide-containing sanitizer against single species and binary
biofilms of a dairy-associated Bacillus cereus and a Pseudomonas fluorescens
isolate. J Appl Microbiol 92(2):352–61.
Lomander A, Schreuders P, Russek-Cohen E, Ali L. 2004. Evaluation of
chlorines’ impact on biofilms on scratched stainless steel surfaces. Bioresour
Technol 94(3):275–83.
Mafu AA, Roy D, Goulet J, Magny P. 1990. Attachment of Listeria
monocytogenes to stainless steel, glass, polypropylene, and rubber surfaces after
short contact times. J Food Prot 53(9):742–6.
Manz W, Wendt-Potthoff K, Neu TR, Szewzyk U, Lawrence JR. 1999.
Phylogenetic composition, spatial structure, and dynamics of lotic bacterial
biofilms investigated by fluorescent in situ hybridization and confocal laser
scanning microscopy. Microb Ecol 37(4):225–37.
Marchand S, Heylen K, Messens W, Coudijzer K, De Vos P, Dewetinck K,
Herman L, De Block J, Heyndrickx M. 2009a. Seasonal influence on
heat-resistant proteolytic capacity of P. lundensis and P. fragi, predominant
milk spoilers isolated from Belgian raw milk samples. Environ Microbiol
11(2):467–82.
Marchand S, Vandriesche G, Coorevits A, Coudijzer K, De Jonghe V,
Dewettinck K, De Vos P, Devreese B, Heyndrickx M, De Block J. 2009b.
Heterogeneity of heat-resistant proteases from milk Pseudomonas species. Intl
J Food Microbiol 133:68–77
Marouani-Gadri N, Firmesse O, Chassaing D, Sandris-Nielsen D, Arneborg
N, Carpentier B. 2010. Potential of Escherichia coli O157:H7 to persist and
form viable but non-culturable cells on a food-contact surface subjected to
cycles of soiling and chemical treatment. Intl J Food Microbiol
144(1):96–103.
Vol. 11, 2012 r Comprehensive Reviews in Food Science and Food Safety 145
Biofilm risks in dairy practice . . .
MattilaSandholm T, Wirtanen G. 1992. Biofilm formation in the
industry—a review. Food Rev Intl 8(4):573–603.
Mccoy WF, Bryers JD, Robbins J, Costerton JW. 1981. Observations of
fouling biofilm formation. Can J Microbiol 27(9):910–7.
McLandsborough L, Rodriguez A, Perez-Conesa D, Weiss J. 2006. Biofilms:
at the interface between biophysics and microbiology. Food Biophys
1(2):94–114.
Mettler E, Carpentier B. 1998. Variations over time of microbial load and
physicochemical properties of floor materials after cleaning in food industry
premises. J Food Prot 61(1):57–65.
Mittelman MW. 1998. Structure and functional characteristics of bacterial
biofilms in fluid processing operations. J Dairy Sci 81(10):2760–4.
Mittelman MW, Kohring LL, White DC. 1990. The role of biofilms in
contamination of process fluids by biological particulates. In: Particles in
gases and liquids, Vol. II. Mittal, New York: Plenium Press. p 33–50.
Moller S, Sternberg C, Andersen JB, Christensen BB, Ramos JL, Givskov
M, Molin S. 1998. In situ gene expression in mixed-culture biofilms:
evidence of metabolic interactions between community members. Appl
Environ Microbiol 64(2):721–32.
Momba MNB, Kfir R, Venter SN, Cloete TE. 2000. An overview of biofilm
formation in distribution systems and its impact on the deterioration of
water quality. Water Sa 26(1):59–66.
Morgenroth E. 2003. Detachment: an often overlooked phenomenon in
biofilm research and modeling. In: Bishop B, Wuertz S, Wilderer P, editors.
Biofilms in wastewater treatment, an interdisciplinary approach. London,
UK: IWA Publishing House. p 264–93.
Mosteller TM, Bishop JR. 1993. Sanitizer efficacy against attached bacteria
in a milk biofilm. J Food Prot 56(1):34–41.
Munsch-Alatossava P, Alatossava T. 2006. Phenotypic characterization of raw
milk-associated psychrotropic bacteria. Microbiol Res 161:334–46.
Murphy PM, Lynch D, Kelly PM. 1999. Growth of thermophilic spore
forming bacilli in milk during the manufacture of low heat powders. Intl J
Dairy Technol 52(2):45–50.
Mustapha A, Liewen MB. 1989. Destruction of Listeria monocytogenes by
sodium-hypochlorite and quaternary ammonium sanitizers. J Food Prot
52(5):306–11.
Nickel JC, Ruseska I, Wright JB, Costerton JW. 1985. Tobramycin
resistance of Pseudomonas aeruginosa cells growing as a biofilm on urinary
catheter material. Antimicrob Agents Chemother 27(4):619–24.
Norwood DE, Gilmour A. 2000. The growth and resistance to sodium
hypochlorite of Listeria monocytogenes in a steady-state multispecies biofilm.
J Appl Microbiol 88(3):512–20.
O’Toole G, Kaplan HB, Kolter R. 2000. Biofilm formation as microbial
development. Ann Rev Microbiol 54:49–79.
O’Toole GA, Kolter R. 1998. Initiation of biofilm formation in Pseudomonas
fluorescens WCS365 proceeds via multiple, convergent signalling pathways: a
genetic analysis. Mol Microbiol 28(3):449–61.
Okabe S, Itoh T, Satoh H, Watanabe Y. 1999. Analyses of spatial
distributions of sulfate-reducing bacteria and their activity in aerobic
wastewater biofilms. Appl Environ Microbiol 65(11):5107–16.
Oliveira K, Oliveira T, Teixeira P, Azeredo J, Henriques M, Oliveira R.
2006. Comparison of the adhesion ability of different Salmonella enteritidis
serotypes to materials used in kitchens. J Food Prot 69(10):2352–6.
Oliver SP, Jayarao BM, Almeida RA. 2005. Foodborne pathogens in milk
and the dairy farm environment: food safety and public health implications.
Foodborne Pathogens Dis 2(2):115–29.
Oulahal N, Martial-Gros A, Bonneau M, Blum LJ. 2004. Combined effect of
chelating agents and ultrasound on biofilm removal from stainless steel
surfaces. Application to “Escherichia coli milk” and “Staphylococcus aureus
milk” biofilms. Biofilms 1:165–73.
Oulahal-Lagsir N, Martial-Gros A, Bonneau M, Blum LJ. 2000. Ultrasonic
methodology coupled to ATP bioluminescence for the non-invasive
detection of fouling in food processing equipment—validation and
application to a dairy factory. J Appl Microbiol 89(3):433–41.
Palmer JS, Flint SH, Schmid J, Brooks JD. 2010. The role of surface charge
and hydrophobicity in the attachment of Anoxybacillus flavithermus isolated
from milk powder. J Ind Microbiol Biotechnol 37(11):1111–9.
Parkar SG, Flint SH, Brooks JD. 2004. Evaluation of the effect of cleaning
regimes on biofilms of thermophilic bacilli on stainless steel. J Appl
Microbiol 96(1):110–6.
Peng JS, Tsai WC, Chou CC. 2001. Surface characteristics of Bacillus cereus
and its adhesion to stainless steel. Intl J Food Microbiol 65(1–2):105–11.
Pereira A, Melo LF. 2009. Monitoring of biofilms in the food and beverage
industries. In: Fratamico PM, Annous BA, Gunther NW, IV, editors.
Biofilms in the food and beverage industries. Oxford, Cambridge, New
Delhi: Wood head Publishing Limited. p 131–51.
Pereira MO, Kuehn M, Wuertz S, Neu T, Melo LF. 2002. Effect of flow
regime on the architecture of a Pseudomonas fluorescens biofilm. Biotechnol
Bioeng 78(2):164–71.
Rice SA, Koh KS, Queck SY, Labbate M, Lam KW, Kjelleberg S. 2005.
Biofilm formation and sloughing in Serratia marcescens are controlled by
quorum sensing and nutrient cues. J Bacteriol 187(10):3477–85.
Rijnaarts HHM, Norde W, Bouwer EJ, Lyklema J, Zehnder AJB. 1993.
Bacterial adhesion under static and dynamic conditions. Appl Environ
Microbiol 59(10):3255–65.
Ruckert A, Ronimus RS, Morgan HW. 2004. A RAPD-based survey of
thermophilic bacilli in milk powders from different countries. Intl J Food
Microbiol 96(3):263–72.
Rysstad G, Kolstad J. 2006. Extended shelf life milk—advances in
technology. Intl J Dairy Technol 59(2):85–96.
Ryu JH, Beuchat LR. 2005. Biofilm formation and sporulation by Bacillus
cereus on a stainless steel surface and subsequent resistance of vegetative cells
and spores to chlorine, chlorine dioxide, and a peroxyacetic acid-based
sanitizer. J Food Prot 68(12):2614–22.
Sasahara KC, Zottola EA. 1993. Biofilm formation by Listeria monocytogenes
utilizes a primary colonizing microorganism in flowing systems. J Food Prot
56(12):1022–8.
Sauer K, Rickard AH, Davies DG. 2007. Biofilms and biocomplexity.
Microbe 2(7): 347–55.
Schwan WR, Seifert HS, Duncan JL. 1992. Growth conditions mediate
differential transcription of fim genes involved in phase variation of type 1
pili. J Bacteriol 174:2367–75.
Scott SA, Brooks JD, Rakonjac J, Walker KMR, Flint SH. 2007. The
formation of thermophilic spores during the manufacture of whole milk
powder. Intl J Dairy Technol 60(2):109–17.
Shah NP. 1994. Psychrotrophs in milk: a review. Milchwissenschaft
49(8):432–7.
Shah NP. 2000. Effects of milk-derived bioactives: an overview. Br J Nutr
84(Suppl 1):S3–S10.
Shaheen R, Svensson B, Andersson MA, Christiansson A, Salkinoja-Salonen
M. 2010. Persistence strategies of Bacillus cereus spores isolated from dairy silo
tanks. Food Microbiol 27(3):347–55.
Sheng XX, Ting YP, Pehkonen SO. 2008. The influence of ionic strength,
nutrients and pH on bacterial adhesion to metals. J Colloid Interface Sci
321(2):256–64.
Sigua G, Adhikari S, Frankel GS, Pascall MA. 2010. The use of atomic force
microscopy to measure the efficacies of various chemical sanitizers in
removing organic matter from glass surfaces. J Food Eng 100(1):139–44.
Silva S, Teixeira P, Oliveira R, Azeredo J. 2008. Adhesion to and
viability of Listeria monocytogenes on food contact surfaces. J Food Prot 71(7):
1379–85.
Simões M, Simões LC, Vieira MJ. 2008. Physiology and behavior of
Pseudomonas fluorescens single and dual strain biofilms under diverse
hydrodynamics stresses. Intl J Food Microbiol 128(2):309–16.
Simoes M, Simoes LC, Vieira MJ. 2009. Species association increases biofilm
resistance to chemical and mechanical treatments. Water Res 43(1):229–
37.
Sirianuntapiboon S, Jeeyachok N, Larplai R. 2005. Sequencing batch reactor
biofilm system for treatment of milk industry wastewater. J Environ Manage
76(2):177–83.
Somers EB, Johnson ME, Wong ACL. 2001. Biofilm formation and
contamination of cheese by nonstarter lactic acid bacteria in the dairy
environment. J Dairy Sci 84(9):1926–36.
Sommer P, Martin-Rouas C, Mettler E. 1999. Influence of the adherent
population level on biofilm population, structure and resistance to
chlorination. Food Microbiol 16(5):503–15.
Sorhaug T, Stepaniak L. 1997. Psychrotrophs and their enzymes in milk and
dairy products: quality aspects. Trends Food Sci Technol 8:35–41.
Speers JGS, Gilmour A. 1985. The influence of milk and milk components
on the attachment of bacteria to farm dairy equipment surfaces. J Appl
Bacteriol 59(4):325–32.
Sreekumari KR, Sato Y, Kikuchi Y. 2005. Antibacterial metals—a viable
solution for bacterial attachment and microbiologically influenced
corrosion. Mater Trans 46(7):1636–45.
146 Comprehensive Reviews in Food Science and Food Safety r Vol. 11, 2012
c 2012 Institute of Food Technologists®
Biofilm risks in dairy practice . . .
Stadhouders J, Hup G, Hassing F. 1982. The conceptions index and
indicator organisms discussed on the basis of the bacteriology of spray-dried
milk powder. Neth Milk Dairy J 36(3):231–60.
Stepanovic S, Cirkovic I, Ranin L, Svabic-Vlahovic M. 2004. Biofilm
formation by Salmonella spp. and Listeria monocytogenes on plastic surface.
Lett Appl Microbiol 38(5):428–32.
Stewart JC, Seiberling DA. 1996. Clean in place. Chem Eng 103(1):
72–9.
Stickler D, Hewett P. 1991. Activity of antiseptics against biofilms of mixed
bacterial species growing on silicone surfaces. Eur J Clin Microbiol Infect
Dis 10(5):416–21.
Stoodley P, Sauer K, Davies DG, Costerton JW. 2002. Biofilms as complex
differentiated communities. Annu Rev Microbiol 56:187–209.
Storgards E, Simola H, Sjoberg AM, Wirtanen G. 1999a. Hygiene of gasket
materials used in food processing equipment part 1: new materials. Food
Bioproducts Process 77(C2):137–45.
Storgards E, Simola H, Sjoberg AM, Wirtanen G. 1999b. Hygiene of gasket
materials used in food processing equipment part 2: aged materials. Food
Bioproducts Process 77(C2):146–55.
Suarez B, Ferreiros CM, Criado MT. 1992. Adherence of psychrotrophic
bacteria to dairy equipment surfaces. J Dairy Res 59(3):381–8.
Sundheim G, Hagtvedt T, Dainty R. 1992. Resistance of meat associated
Staphylococci to a quarternary ammonium compound. Food Microbiol
9(2):161–7.
Tamachkiarow A, Flemming HC. 2003. On-line monitoring of biofilm
formation in a brewery water pipeline system with a fibre optical device.
Water Sci Technol 47(5):19–24.
Tang XM. 2011. Controlling biofilm development on ultrafiltration and
reverse osmosis membranes used in dairy plants, Thesis submitted in
fulfilment of the requirements of the degree of Doctor in Phylosophy in
Food Technology, Massey Univ., Manawatu, New Zealand, p 1–152.
Tang XM, Flint SH, Bennett RJ, Brooks JD. 2010. The efficacy of different
cleaners and sanitisers in cleaning biofilms on UF membranes used in the
dairy industry. J Membr Sci 352(1–2):71–5.
Tang XM, Flint SH, Brooks JD, Bennett RJ. 2009a. Factors affecting the
attachment of micro-organisms isolated from ultrafiltration and reverse
osmosis membranes in dairy processing plants. J Appl Microbiol
107(2):443–51.
Tang XM, Flint SH, Bennett RJ, Brooks JD, Morton RH. 2009b. Biofilm
growth of individual and dual strains of Klebsiella oxytoca from the dairy
industry on ultrafiltration membranes. J Ind Microbiol Biotechnol
36(12):1491–7.
Tauveron G, Slomianny C, Henry C, Faille C. 2006. Variability among
Bacillus cereus strains in spore surface properties and influence on their ability
to contaminate food surface equipment. Intl J Food Microbiol
110(3):254–62.
te Giffel MC, Beumer RR, Bonestroo MH, Rombouts FM. 1996a.
Incidence and characterization of Bacillus cereus in two dairy processing
plants. Neth Milk Dairy J 50:479–92.
te Giffel MC, Beumer RR, Leijendekkers S, Rombouts FM. 1996b.
Incidence of Bacillus cereus and bacilus subtilis in foods in the Netherlands.
Food Microbiol 13:53–8.
Teixeira P, Lopes Z, Azeredo J, Oliveira R, Vieira MJ. 2005. Physico-chem-
c 2012 Institute of Food Technologists®
ical surface characterization of a bacterial population isolated from a milking
machine. Food Microbiol 22(2–3):247–51.
Telgmann U, Horn H, Morgenroth E. 2004. Influence of growth history
on sloughing and erosion from biofilms. Water Res 38(17):3671–84.
Ternström A, Lindberg A-M, Molin G. 1993. Classification of the spoilage
flora of raw and pasteurized bovine milk, with special reference to
Pseudomonas and Bacillus. J Appl Bacteriol 75:25–34.
Trachoo N. 2003. Biofilm and food industry. J Sci Technol 25:807–15.
van den Broek D, Bloemberg GV, Lugtenberg B. 2005a. The role of
phenotypic variation in rhizosphere Pseudomonas bacteria. Environ
Microbiol 7(11):1686–97.
van den Broek D, Chin AW, Bloemberg GV, Lugtenberg BJJ. 2005b.
Molecular nature of spontaneous modifications in gacS which cause colony
phase variation in Pseudomonas sp strain PCL1171. J Bacteriol
187(2):593–600.
Vanloosdrecht MCM, Lyklema J, Norde W, Zehnder AJB. 1989. Bacterial
adhesion—a physicochemical approach. Microb Ecol 17(1):1–15.
Werner E, Roe F, Bugnicourt A, Franklin MJ, Heydorn A, Molin S, Pitts B,
Stewart PS. 2004. Stratified growth in Pseudomonas aeruginosa biofilms. Appl
Environ Microbiol 70(10):6188–96.
Whitchurch CB, Tolker-Nielsen T, Ragas PC, Mattick JS. 2002.
Extracellular DNA required for bacterial biofilm formation. Science
295(5559):1487.
White-Ziegler CA, Black AM, Eliades SH, Young S, Porter K. 2002.
The N-acetyltransferase Rimj responds to enviromental stimuli to
repress pap fimbrial transcription in Escherichia coli. J Bacteriol 184:
4334–42.
White-Ziegler CA, Villapakkam A, Ronaszeki K, Young S. 2000. H-NS
control pap and daa fimbrial transcription in Escherichia coli in response to
multiple environmental cues. J Bacteriol 182:6391–400.
Wijman JGE, de Leeuw PPLA, Moezelaar R, Zwietering MH, Abee T.
2007. Air-liquid interface biofilms of Bacillus cereus: formation, sporulation,
and dispersion. Appl Environ Microbiol 73(5):1481–8.
Wimpenny J, Manz W, Szewzyk U. 2000. Heterogeneity in biofilms. Fems
Microbiol Rev 24(5):661–71.
Wingender J, Strathmann M, Rode A, Leis A, Flemming HC. 2001.
Isolation and biochemical characterization of extracellular polymeric
substances from Pseudomonas aeruginosa. Microb Growth Biofilms, A 336:
302–14.
Wirtanen G. 1995. Biofilm formation and its elimination from food
processing equipment, Vol. 251. Espoo: VTT Technical Research Centre of
Finland.
Wirtanen G. 2004. Hygiene control in Nordic dairies, Vol. 545. ESPOO:
VTT Publication. p 147–62.
Wong ACL. 1998. Biofilms in food processing environments. J Dairy Sci
81(10):2765–70.
Wong ACL, Cerf O. 1995. Biofilms: implications for hygiene monitoring of
dairy plant surfaces. Bull IDF 302:40–4.
Workentine ML, Harrison JJ, Weljie AM, Tran VA, Stenroos PU, Tremaroli
V, Vogel HJ, Ceri H, Turner RJ. 2010. Phenotypic and metabolic profiling
of colony morphology variants evolved from Pseudomonas fluorescens
biofilms. Environ Microbiol 12(6):1565–77.
Vol. 11, 2012 r Comprehensive Reviews in Food Science and Food Safety 147