Biological Aspects in Food Preservation by Ultraviolet ...ssu.ac.ir/cms/fileadmin/user_upload/Mtahghighat/... · impact of these factors on microbial UV resistance must be well understood

REVIEW

Biological Aspects in Food Preservation by UltravioletLight: a Review

Elisa Gayán & Santiago Condón & Ignacio Álvarez

Received: 15 February 2013 /Accepted: 29 July 2013 /Published online: 17 August 2013# Springer Science+Business Media New York 2013

Abstract The potential to commercialize nonthermal ultravi-olet (UV) light technologies as new methods for preservingfood products has caught the attention of a food industry thatwishes to fulfill consumers' demands for fresh products.Numerous investigations have demonstrated UV light's abilityto inactivate a wide range of microorganisms. However, thelack of UV sensitivity data from pathogenic and spoilagebacteria is evident. In addition, the main factors associatedwith UV light in terms of microbial lethality remain unclear.This review surveys critical factors (process, microbial, andenvironmental parameters) that determine UV microbial resis-tance and assess the effects of such factors on the inactivationmechanism and repair pathway efficiency. The effects of someof these factors, such as prior sublethal stresses and post-recovery conditions of UV treatments, may extensively im-prove the damage repair capacity and thus microbial surviv-ability. Further research is needed to establish adequate controlmeasures pre- and post-UV treatments. Furthermore, the pos-sibility of combining UV light with conventional preservativesand other nonthermal technologies was assessed. The combi-nation of UV light with mild heating or oxidant compoundscould offer promising treatments to enhance the safety andstability of minimally processed foods.

Keywords UV light . Bacteria inactivation . DNA repair .

Sub-lethal stress . Combined processes

Abbreviations6-4PP Pyrimidine 6–4 pyrimidone photoproduct8-oxodGuo 8-Oxo-7,8-dihydro-2′-deoxyguanosineAOP Advanced oxidation processesBER Base excision repair

CPD Cyclobutan pyrimidine dimerDewPP Dewar valence isomersDSB Double-strand breakHS Heat shockHSP Heat shock proteinsIR Infrared radiationLHR Liquid holding recoveryLP Low-pressure mercury vapor lampsMM Minimal mediumMP Medium-pressure mercury vapor lampsNER Nucleotide excision repairppGpp Guanosine 5′-diphosphate 3′-diphosphatePRR Post-replication repairPUV Pulsed UV lampPX Pulsed xenon lampRAMER RecA-mediated excision repairROS Reactive oxygen speciesSP Spore photoproductSSB Single-strand breakTLS Translesion DNA synthesis

Introduction

Traditionally, ultraviolet (UV) light irradiation has been usedas a disinfectant for air, surface, and water decontamination.Recently, the food industry has shown increasing interest inusing UV irradiation for the hygienization of liquid foods andthe surfaces of solid foods. As an emerging nonthermal pro-cessing technology, UV irradiation offers multiple advantages:effective inactivation of a broad range of spoilage and patho-genic microorganisms, minimal loss of the nutritional andsensorial quality of foods, no known toxic effects or residuesfrom the treatment, and low energy consumption compared toother thermal and nonthermal pasteurization processes.

E. Gayán : S. Condón : I. Álvarez (*)Tecnología de los Alimentos, Facultad deVeterinaria, Universidad deZaragoza, C/ Miguel Servet 177, CP 50013 Zaragoza, Spaine-mail: [email protected]

Food Bioprocess Technol (2014) 7:1–20DOI 10.1007/s11947-013-1168-7

UV light refers to the part of the electromagnetic spectrumwhich ranges from 200 to 400 nm, and it is divided into threeregions: short-wave ultraviolet (UV-C), from 200 to 280 nm;UV-mediumwave (UV-B), from 280 to 320 nm; and UV-longwave (UV-A), from 320 to 400 nm. The effects of UV light ongenetic material are primarily responsible for microbial inac-tivation, although other cellular components such as proteinscan be also damaged. UV-C constitutes the most germicidalregion, and the peak of maximum effectiveness is at wave-lengths of about 260–265 nm, corresponding with the peak ofmaximum DNA absorption (Fig. 1) (Kowalski 2009). UV-Cabsorption induces the formation of DNA photoproducts, inparticular cyclobutane pyrimidine dimers and pyrimidine 6–4pyrimidone photoproducts, which inhibit transcription andreplication and eventually lead to mutagenesis and cell death.

Since UV-related microbial inactivation arises primarilyfrom the effects of UV light on genetic material, the UVresistance of a specific microorganism depends on the effec-tiveness of its DNA repair mechanisms as well as the extent ofDNA damage induced by the treatment (López-Malo andPalou 2005). In addition, DNA damage and repair mecha-nisms are affected by different factors, classified as microbial,environmental, and processing factors, which can act before,during, or after UV treatments.

For instance, bacteria change their physiology under differ-ent growing conditions (growth medium, incubation tempera-ture, and growth rate), which could vary their UV resistancesignificantly (López-Malo and Palou 2005). Stress historyprior to UV treatment, such as heat, acid, osmotic, and starva-tion shocks, may also influence resistance by triggering co-protective adaptation responses (Van der Veen and Abee2011). Recovery conditions also affect cell viability after irra-diation, especially the exposure to visible light and growth orrecovery medium composition (Ganesan and Smith 1968;Sommer et al. 2000). Temperature increments during treatmentand the addition of prooxidant chemical compounds mayimprove UV lethality (Koivunen Heinonen-Tanski 2005;

Gayán et al. 2011). Consequently, as UV light technologygains interest as a strategy for food decontamination, theimpact of these factors on microbial UV resistance must bewell understood in order to assess the actual lethality implica-tions of UV light.

Thus, the objective of this review is to compile up-to-dateknowledge about the influence of process, microbial, andenvironmental factors that could affect UV treatments forbacteria and bacterial spores. This review will evaluate theinfluence of these factors on UV survivability and lethality onthe basis of inactivation and damage repair. To achieve thisgoal, the review will first describe the general mechanisms ofinactivation and repair. Based on those, the influence of thelethality and survivability factors will be discussed.

Mechanisms of Inactivation by UV Radiation

Genetic materials are the primary targets of UV radiation, andthe types of lesions produced depend on the radiation band(UV-C, UV-B, or UV-A). UV-C induces the formation ofphotoproducts due to the direct absorption of photons bypyrimidine and purine nucleic acid bases (López-Malo andPalou 2005). Although both pyrimidine and purine bases arestrong UV-C photon absorbers, pyrimidines absorb about 10times more than purines. Therefore, photoproducts derivedfrom pyrimidines are the most important (Kowalski 2009).The major lesions induced by UV-C are cyclobutane pyrimi-dine dimers (CPDs) and pyrimidine 6–4 pyrimidone photo-products (6-4PPs) (Friedberg et al. 2006).

CPDs are formed when there are two adjacent pyrimidinesand a photon being absorbed by one pyrimidine. Pyrimidinesbecome covalently linked when the saturation of the carbonatoms (C) 5 and 6 of both neighboring bases causes theformation of a ring structure (Fig. 2(a)). CPDs are formedbetween thymines (TT), cytosines (CC), or cytosine and thy-mine (CTor TC) bases, depending on nucleotide composition.In general, dimers containing thymines are most likely tooccur (Rastogi et al. 2010).

A similar process creates 6-4PPs, except that adducts aregenerated by cycling between the C5–C6 bond of a pyrimi-dine and the C4 carbonyl or imino groups (for thymine orcytosine, respectively) for its 3′ neighbor. The resultingoxetane and azetidine products are unstable and rapidlyrearrange themselves, transferring the carbonyl or aminogroup of the 3′ base to the C5 position of the 5′ base, yieldingthe 6-4PP (Fig. 2(c)). In this case, 6-4PPs are more frequentlyobserved with CC and TC, while TT or CT sequences are lesscommon. When exposed to UV-A or UV-B, 6-4PP can beconverted into Dewar valence isomers (DewPPs). However,this damage is less common (Rastogi et al. 2010).

The proportion of 6-4PPs is much lower than that of CPDs,accounting for about 25 % of total UV-mediated DNA

Fig. 1 Comparisons of the relative germicidal efficiency of UV wave-lengths, comparing LP (1), MP (2), and PX lamps (3) with germicidaleffectiveness for E. coli (dotted line)

2 Food Bioprocess Technol (2014) 7:1–20

damage (Sinha and Hader 2002). This is probably because theefficiency of the dimerization process, given by the quantumyield (Φ), is higher for CPDs (Φ254 nm=22–26) than for 6-4PPs (Φ254 nm=1.4) (Görner 1994). However, 6-4PPs arepotentially more lethal and mutagenic than CPDs because oflower repair efficiency (Koehler et al. 1996). Both CPDs and6-4PPs cause structural distortions in DNA molecules thatinterfere with normal RNA transcription and DNA replication,leading to mutagenesis and ultimately to cell death (Friedberget al. 2006).

Bacterial spores are also affected when they are exposed toUV-C light. Spore photoproducts (SPs) are the major DNAlesions in UV-irradiated bacterial spores (Douki et al. 2005;Moeller et al. 2007). SPs are formed by the addition of amethyl group of thymine residue to the C5 position of anadjacent thymine (5-thyminyl-5,6-dihydrothymine, Fig. 2(b))(Rastogi et al. 2010). SP formation is related to the dehydratedA-conformation of the DNA molecule. Hence, SPs have alsobeen isolated in vegetative cells at low relative humidity(Peccia et al. 2001). The consequences of SPs are similar tothose indicated for CPDs and 6-4PPs.

CPDs and 6-4PPs are also produced by bacterial spores, butin very small quantities compared to SPs (Douki et al. 2005;Moeller et al. 2007). For instance, Douki et al. (2005) reportedthat the CPD yield induced by UV-C in Bacillus subtilis sporeswas only 0.1 % that of SPs, and the levels for 6-4PPs wereeven lower. This could be attributed to the fact that the quan-tum efficiency for SP formation by 254 nm radiation (252

lesions per 104 per J/cm2) is higher than for the generation ofother photoproducts (1.0 CPD and 0.1 6-4PP per 104 per J/cm2) (Setlow 2006). However, SPs are much less lethal thanCPDs or 6-4PPs, as SPs can be repaired efficiently in the firstminutes of spore germination via different mechanisms(Setlow 2006). This could also explain the higher UV-C resis-tance of bacterial spores compared to their correspondingvegetative cells (Setlow 2001), since the quantum yield forSP formation in bacterial spores is similar to that for CPD and6-4PP in growing cells (Setlow 2006). Single- and double-strand breaks (SSBs and DSBs) can be also detected at highUV fluence to a small extent. Slieman and Nicholson (2000)applied a UV-C dose of 16 kJ/m2 to dried B. subtilis spores.Whereas nearly 40 % of the total chromosomal thymine wasconverted within the SPs, only 0.3 and 0.16 % were present inthe SSBs and CPDs, respectively. Thus, CPDs, SSBs, andDSBs probably do not have major physiological consequencesfor the UV-C survival of spores (Slieman andNicholson 2000).

When cells are exposed to UV-B radiation, fewer CPDs, 6-4PPs, and DewPPs are produced compared to those in UV-Cradiation (Ravanat et al. 2001; Pattison andDavies 2006). Otherphotoproducts may occur to a smaller extent, such as cytosinephotohydration, guanine bases oxidations, or adduct formationbetween either two adjacent adenine bases or between adenineand vicinal thymine (reviewed in Ravanat et al. 2001).

UV-A radiation produces less photochemical damage be-cause of DNA's low absorbability in this range. CPDs areformed about 105-fold less efficiently than at 254 nm, and 6-

Fig. 2 Primary DNA photoproducts caused by UV-C and UV-B: cyclobutan pyrimidine dimer (CPD) (a), spore photoproduct (SP) (b), pyrimidine 6–4pyrimidone photoproduct (6-4PP) (c), and 8-oxo-7,8-dihydro-2′-deoxyguanosine (8-oxodGuo) (d)

Food Bioprocess Technol (2014) 7:1–20 3

4PPs are barely detectable. On the other hand, DewPPs areinduced more frequently here than by UV-B (Girard et al.2011). DNA photooxidation damage of pyrimidine and purinebases is also produced by UV-A. Guanine is the base mostsusceptible to oxidization, resulting in 8-oxo-7,8-dihydro-2′-deoxyguanosine (8-oxodGuo, Fig. 2(d)), the most frequentlyseen oxidative lesion under UV-A radiation (Douki et al.2003). Oxidation-damaged bases result in nonbulky lesionsthat exert fewer obstacles for replication and transcription,leading to DNA mutagenesis and the production of mutantproteins (transcriptional mutagenesis). In addition, UV-A ra-diation can also produce SSBs, but in very low numbers.Overall, despite the low energy of UV-A photons for pyrim-idine dimer formation, CPDs represent the main source ofDNA damage, followed by purine oxidations (essentially 8-oxoG), pyrimidine oxidations, and SSBs in a ratio of 10:3:1:1(Douki et al. 2003).

Finally, besides DNA damage, UV-A photooxidation af-fects other biological targets such as proteins, lipids, andsterols. In fact, proteins are major cellular targets for photo-oxidation due to both their high abundance and the presenceof endogenous chromophores within their structures (certainamino acid side chains, as well as bound chromophores suchas flavins and porphyrins). Direct damage induced by UV-Aradiation is limited to a small number of amino acid residues,mainly tryptophan (Trp), tyrosine (Tyr), histidine (His), anddisulfide (cystine). However, most UV-induced protein dam-age occurs indirectly, mediated by reactive oxygen species(ROS) which react preferentially with Trp, His, Tyr, methio-nine (Met), cysteine (Cys), and cystine side chain residues.When these residues are initially altered, multiple secondaryprocesses occur, resulting in damage to other intra- andintermolecular sites. These processes can lead to protein frag-mentation, aggregation, and altered physical and chemicalproperties, resulting in the loss of functionality (reviewed inPattison and Davies 2006). UV-A can inactivate DNA repairenzymes such as photolyase (Tyrrell et al. 1973) andglycosylases (Eiberger et al. 2008). However, the main causeof cell death is DNA molecule alteration.

As observed, there are many forms of UV-induced damagedetermining the microbial lethality of UV treatments.Nevertheless, microbial survivability against UV light is alsodependent on the cells' capacity to repair damage. As there aremany different types of UV-induced DNA damage, microor-ganisms have developed several pathways for DNA repair.

DNA Repair Mechanisms

DNA repair pathways are classified according to temporalintervention (prior, during, or after replication) and actionmechanism. Three main pathways are involved in DNA re-pair: (1) reverse damage repair and (2) excision repair

systems, which act before cell replication occurs, and (3)tolerating damage pathways, which are activated once repli-cation has started or immediately afterward. The main DNArepair pathways are briefly described below and presented inFig. 3.

The first group of repair mechanisms restores the DNAmolecule before replication, with remarkable precision andspeed. However, when DNA damage remains unrepaired,DNA polymerase III (Pol III), the enzyme responsible ofDNA replication, is blocked, and the SOS response to DNAdamage is activated (Bichara et al. 2011). The SOS response isa global regulatory network governed by the nucleofilamentRecA (comprising RecA proteins and single-stranded DNA),which stimulates cleavage of the LexA repressor. This inducesthe transcription of more than 40 genes involved in celldivision delay, alternative repair mechanisms, and toleratinglesion pathways, permitting replication (Janion 2008).

Reverse damage mechanisms are carried out by DNAlyases, which restore the damaged part of the molecule in situ,without DNA synthesis. In general, two kinds of DNA lyasescan be distinguished in bacteria: CPD lyases (also calledphotolyases in the mechanism known as “photorepair”) andSP lyase. CPD lyases are enzymes responsible for photoreac-tivation in vegetative bacteria cells, acting specifically againstCPDs using the energy of visible light (350–500 nm). That is,visible light is necessary for them to act (Sinha and Hader2002). SP lyase is the main enzyme of SP repair in spores,acting during spore germination (Setlow 2006). It does notrequire visible light. Besides the SP lyase mechanism, SP canbe removed by excision repair or RecA-mediated pathways,which play minor roles in germinating spore repair (Moelleret al. 2007).

Excision repair mechanisms replace damaged DNA byresynthesizing nucleotides. They are considered error-freemechanisms because excised nucleotides are replaced byDNA polymerase I (Pol I) using the parental strand as atemplate. There are two types of excision repair: base excisionrepair (BER), which first removes the damaged base, leavingan apurinic/apyrimidinic site that is then removed; and nucle-otide excision repair (NER), which directly removes the nu-cleotide tracts containing lesions. The key enzymes involvedin BER are DNA glycosylases that recognize specific lesionsin nitrogen bases and excise them. An example of DNAglycosylase is the endonuclease Vor pyrimidine dimer DNAglycosylase, normally present in UV-resistant microorganismssuch as Micrococcus luteus, Escherichia coli phage T4, andSaccharomyces cerevisiae (Jacobs and Schaer 2012). NERcan remove a broad spectrum of DNA lesions through multi-ple cascade reactions carried out primarily by the UvrABCexinuclease (reviewed in Truglio et al. 2006). NER carried outby constitutive UvrABC complex is also referred to as “darkrepair” or “liquid holding recovery” (LHR) because UV-irradiated cells are able to repair their DNA suspended in


minimal medium without the presence of light (Ganesan andSmith 1968).

DNA damage tolerance and repair mechanisms inducedafter replication activation include RecA-mediated excisionrepair (RAMER), translesion DNA synthesis (TLS), and post-replication repair (PRR) (reviewed in Bichara et al. 2011). Theactivities of all of these mechanisms are regulated thoroughlyin the temporal space by the SOS regulon to safeguard ge-nome integrity.

One of the first such mechanisms to be induced is NERmediated by the action of RecA protein (RAMER). Whenthere is extensive damage, the expression of UvrABCexinuclease genes under SOS regulon control (uvrA anduvrB) is reinforced. This provides an effective second oppor-tunity for NER to remove lesions, with the help of Rec familyproteins (Courcelle et al. 1999).

TLS occurs during replication and consists of the misin-corporation of nucleotides into the synthesized strand oppositethe lesion by specialized polymerases, giving rise to mutations(Sinha and Hader 2002). Once the lesion is bypassed, Pol IIIregains control of the replication. TLS is carried out primarilyby three SOS-inducible DNA polymerases. The first twoare Pol II and Pol IV, which are induced at the early stagesof the SOS response due to their high error-free activity(nonmutagenic TLS). The third is Pol V (UmuDC complex),which has a high mutagenic activity (mutagenic TLS) and,consequently, low error-free activity. Pol V is activated in thelatest stages of the SOS response as a last resort to recover cellviability (Bichara et al. 2011).

Finally, when a lesion persists, the blockage of Pol III maylead to the formation of gaps or discontinuities in the daughterstrand. These gaps are filled after replication by homologous

T-T

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RE

PL

ICA

TIO

ND

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ING

AN

D P

OS

T R

EP

LIC

AT

ION

Photorepair CPD photolyase

Visible light 350-500 nm

Excision repair

UvrB

UvrA

UvrB

UvrA

UvrC

Pol I

NER

BER

DNA glycosylase

AP site

AP lyase

Pol II, Pol IV (non-mutagenicTLS)

RAMER

SOS regulon

RecA

LexA

RNAp

TLS

Pol III

SOS response

Pol V (mutagenicTLS)

Pol III

PRR

Homologous recombination

CPD

Reverse damage

Pol III

Fig. 3 DNA repair mechanisms in vegetative cells


recombination, a process which is also mainly controlled byRecA (Sinha and Hader 2002). This mechanism has beenreferred to as PRR.

Generally, the action of error-free excision repair pathwayspredominates over tolerating mechanisms to avoid their po-tentially detrimental effects and thereby maintain genetic sta-bility (Bichara et al. 2011). For instance, Bichara et al. (2007)reported that whenDNA lesions were induced inE. coli by thechemical carcinogen N-2-acetylaminofluorene, 80 % of thesurviving colonies benefited from RAMER activity. Theremaining colonies (20 %) survived through homologousrecombination mechanisms, predominating over mutagenesisTLS.

Thus, many mechanisms can act to repair UV-inducedDNA damage. The effects of these mechanisms on microbialsurvivability after UV light treatment vary depending on re-covery conditions (“Recovery Conditions After Processing”section) and could affect microbial resistance and the kineticsof inactivation to a great extent. For example, after UV treat-ment, it has been observed that 99.99 % of the damagedpopulation of some strains of E. coli can recover in the pres-ence of visible light (2 mW/cm2 at 360 nm for 3 h) (Kashimadaet al. 1996). Therefore, it is essential to identify the mainparameters affectingUVresistance and how damage and repairmechanisms may influence microbial resistance to UV light.

Factors Influencing Microbial Resistance to UV Light

Several factors determine microbial inactivation through UVlight. These can be essentially classified as process parame-ters, microbial characteristics, and product parameters(Table 1). An adequate knowledge of these factors and theirinfluence on bacterial death or survivability is necessary toobtain quality UV inactivation data, as well as to preventexperimental artifacts and avoid misinterpreting results.

Process Parameters

The main process parameters affecting microbial resistance toUV light (Table 1) are wavelength emission and UV dose.Other physical factors could affect microbial resistance, suchas conformation and geometry of the UV equipment, flowpattern, and optical characteristics of the treatment medium(López-Malo and Palou 2005). However, these parameters donot have a direct biological effect on microorganisms, only onthe UV dose received by them. This could result in methodo-logical artifacts interfering with the correct determination ofmicrobial inactivation kinetic parameters.

UVeffectiveness on microbial inactivation is dependent onthe wavelength emission. Higher lethal efficacy is achieved atwavelengths closer to DNA's absorption peak. The wavelengthemission depends on the UV lamp used to apply the treatment.

Most conventional UV lamps used for hygienization in thefood and water industries are low-pressure (LP) and medium-pressure (MP) mercury vapor lamps and pulsed xenon (PX)lamps, technology known as pulsed UV light (PUV)(Kowalski 2009).

LP lamps emit quasi-monochromatic light at a wavelengthof 253.7 nm (85 % of total lamp emission), close to maximumDNA absorbance. MP lamps emit a polychromatic spectrumcovering wavelengths from around 200 to 600 nm, with only15–23 % of emissions at 253.7 nm (Fig. 1) (Kowalski 2009).MP lamps show the same microbial survival rate as LP lampsat the same dose, despite the expected damage to other cellularcomponents because of their polychromatic nature (Ogumaet al. 2004; Eischeid and Linden 2007). For instance, Ogumaet al. (2004) reported that the UV doses required to inactivate90 % of the initial population of E. coli and Legionellapneumophila suspended in a phosphate buffer (pH 7.2) usingtwo LP lamps (20 W) and a MP lamp (330 W) were notsignificantly different.

PUV technology has been considered a more rapid andeffective method of microbial inactivation than continuousUV irradiation due to the use of intense, short durationpulses across a broad spectrum (100–1,100 nm) (Fig. 1).The main cause of PUV inactivation is also believed to beDNA damage caused by UV wavelengths (approximately54 % of emitted energy), although concomitant mechanismsmay contribute to improving its effectiveness (Demirci andKrishnamurthy 2011). In addition to photochemical DNAdamage, photothermal effects and photophysical effects havebeen also described. Photothermal effects are attributed to thetemporary intracellular overheating caused by the absorptionof photons, especially from UV-A and UV-B, when highfluence is applied (Wekhof et al. 2001). Wekhof (2000) mea-sured the temperature of E. coli cells treated by different PUVtreatments. Applying a fluence of 0.5 J/cm2, cell temperaturemomentarily rose, reaching values higher than 120 °C.Photophysical effects refer to the disruption of cell membranesand other cell components. The origin of these alterations isnot clear, but some authors have suggested that they areconsequences of temperature increases in bacteria and sur-rounding media (Wekhof et al. 2001; Takeshita et al. 2003).More investigation is needed to elucidate PUV technology'sactual mechanism of microbial inactivation.

The colony-forming viability of bacteria, as well as thetotal DNA damage, is highly correlated with the applied UVdose (Oguma et al. 2004; Eischeid and Linden 2007).However, the inactivation kinetics described in the literaturedo not always follow the same pattern when the microbialinactivation, expressed in logarithmic units, is representedagainst the UV dose (survival curve). Some authors (Oteizaet al. 2010) have described the UV survival curves using first-order kinetics, where the number of viable cells decreasesexponentially when a constant dose is applied:


Ln N=N0ð Þ ¼ e−kd

where N and N0 are the number of microorganisms for anytreatment dose (d) and at the beginning, respectively, and k isthe inactivation rate constant. Considering conventional heattreatments, a decimal reduction dose (DUV) can also be definedas the dose necessary to reduce 90 % of the initial population,and this is equal to 2.303/k. DUV values are commonly used tocharacterize the UV sensitivity of microorganisms.

First-order kinetics is characterized as a one-hit process,assuming that the death of microorganisms is due to a singleevent (the reaction of one UV photon) involving a vital target(DNAmolecule) and that all cells have an identical probabilityof death (Quintero-Ramos et al. 2004). However, in UVsurvival curves, the results often deviate from linearity, show-ing shoulders, tails, or both (Sastry 2000). According to themulti-hit single target theory, the shoulder phase is related todamage and DNA repair phenomena: Various photon events

have to affect the bacterial chromosome until the number oflesions exceeds the capability of DNA repair mechanisms(Quintero-Ramos et al. 2004; López-Malo and Palou 2005).Once these repair mechanisms are overcome, minimal addi-tional UV exposure would be lethal for microorganisms, andthe number of survivors would rapidly decline. This isevidenced by DNA repair-deficient strains, for which survivalcurves show a great reduction in the shoulder phase, accom-panied by an increased slope within the log-linear region(Ganesan and Smith 1968). On the other hand, the upwardconcavity means that the process is becoming progressivelyless efficient with the applied dose. The tailing phase can beexplained by the existence of subpopulations with differentUVresistance and cell aggregates, as well as associations withparticles in the liquid medium (Mamane-Gravetz and Linden2005; Velez-Colmenares et al. 2011).

These biological causes should be distinguished from thoseproduced by the lack of homogeneous dose distribution in thereactor, which is related to the high absorbance or presence of

Table 1 Classification ofinfluencing factors on microbialresistance to UV light. Small let-ters (from a to c) indicate the rel-ative relevance of each factor(a—none, non-influence;b—relevant; c—very relevant)

Processparameters

Microbial characteristics Product parameters

Dose (c) Intrinsic factors pH

Wavelength (c) Type (c) Cell wall thickness awSpecie (b) Cell size Absorption

coefficient (c)

Strain (c) Pigmentation Turbidity (b)

DNA molecule(size, composition,compaction)

Efficiency of repairmechanisms

Membrane fluidity

Extrinsic factors

Growth phase (b)

Growing conditions

Medium composition (a)

Temperature

Stresses

Oxidative shock (a)

Acid shock (b)

Basic shock (a)

Osmotic shock

Heat/IR shock (c)

Starvation stress (b)

Visible light (b)

UV (homologous protection)

Recovery conditions

Medium composition (a)

LHR (a)

Photoreactivation (c)


Table 2 UV doses required for 90 % reduction of the initial population(DUV values) of different types of microorganisms, species, and strainsreported in water and laboratory media. DUV values were expressed influence unit (millijoules per square centimeter), except where indicated.

The type of UV system (bench scale or continuous flow system) and UVsource (number of lamps, type of lamps, and the total input power) usedto obtain data are included

Microorganism Treatment medium UV system UV source DUV (mJ/cm2) Reference

Candida albicans ATCC 10231 Buffer Bench scale LP (17 W) 44.7 Abshire and Dunton (1981)Micrococcus radiodurans ATCC 13939 198.6

Enterococcus faecalis ATCC 10541 12.0

Staphylococcus aureus ATCC 6538 5.6

Escherichia coli ATCC 8739 8.1

Pseudomonas cepacia ATCC 25416 5.2

Pseudomonas diminuta ATCC 11568 11.9

Pseudomonas diminuta ATCC 19146 7.4

Pseudomonas aeruginosaWB-1 5.8

Pseudomonas aeruginosa G-2 3.0

Pseudomonas aeruginosa ATCC 10145 4.6

Acanthamoeba castellanii ATCC 30234 Wastewater NA NA 60 Chang et al. (1985)a

Rotavirus SA-11 7.1

Bacillus subtilis ATCC 6633—spores 36

Enterococcus faecalis ATCC 29212 6.6

Staphylococcus aureus ATCC 25923 3.9


Salmonella typhi ATCC 6539 2.7

Bacillus subtilis DE69 PBS Sunlight MP 3.2 Wassmann et al. (2011)Vegetative cells

Spores Simulator SOL 2 14.2

Escherichia coli O157:H7 CCUG 29193 Buffer Bench scale 10× LP (36 W) 3.5 Sommer et al. (2000)a

Escherichia coli O157:H7 CCUG 29199 0.4

Escherichia coli O25:K98:NM 5


Cryptosporidium parvum Water and wastewater MP 4.4 Hijnen et al. (2006)b

Girdia muris MP 8.2

Acanthamoeba spp. MP 47.6

Bacillus subtilis—spores MP 16.9

Clostridium perfringens—spores LP 16.7

Estreptococcus faecalis MP 3.2

Legionella pneumophila MP 2.5

Escherichia coli MP 1.8

Salmonella typhi MP 1.9

Campylobacter jejuni MP 1.1

Yersinia enterocolitica MP 1.1

Vibrio cholerae MP 0.7

Enterococcus faecium NUIG J952 TSYEA Bench scale PX 4.2c Farrell et al. (2010)Enterococcus faecalis NUIG D46209 3.4c

Enterobacter sakazakii NCTC 8155 2.1c

Staphylococus aureus NUIG 5624 2.0c

Listeria monocytogenes NUIG 11994 2.6c

Escherichia coli DIT 02B1173 2.0c

Klebsiella pneumoniae DIT 04B4415 1.9c

Pseudomonas aeruginosa NUIG 2605 2.7c

Pseudomonas aeruginosa NUIG R5137 1.5c

Pseudomonas aeruginosa NUIG 2633 2.4c


suspended solids in liquid foods, high microbial concentra-tion, and the shading effect from surface irregularities in asolid matrix (Koutchma 2009). For example, in some studiesapplying UV light to surfaces, survival curves with tails havebeen observed (Schenk et al. 2008). It is important to distin-guish tailing phenomena from artifacts resulting from errors inthe methods used for enumerating survivors.

One of the consequences of the absence of linearity in thesurvival curves is the difficulty of obtaining a parameter todescribe inactivation kinetics and compare results. To solve thisproblem, authors have described survival curves with differentmathematic nonlinear equations, such as theWeibull, Gompertz,Geeraerd, and log-logistic models (Quintero-Ramos et al. 2004;Unluturk et al. 2010; Gayán et al. 2011). However, there is noagreement about themost adequatemodel to fit these deviations,and this consensus is necessary to compare results and accurate-ly calculate processing parameters for effective hygienization.

Microbial Characteristics

Microbial resistance to UV light has been found to depend onboth intrinsic and extrinsic factors (Table 1).

Intrinsic Factors

UV resistance varies widely by the type of microorganism,species, and strain. Tables 2 and 3 show some UV resistancedata from the literature based on laboratory media, water, andliquid food to illustrate this variability. Frequently, comparingdata obtained by different authors is risky because of thedifferent conformations and geometry of UV light equipment,flow or stir patterns, and the optical properties of the liquidmedia used, as well as discrepancies in the expression andcalculation of the applied doses. For this reason, methodolog-ical elements used to obtain the resistance data were includedin the tables.

According to Table 2, vegetative bacteria are the mostsensitive, followed by yeast, bacterial spores, viruses, andprotozoa. Inside the bacterial group, Gram-positive bacteriagenerally show more UV resistance than Gram-negatives, butthis is not a general rule. Enterococcus spp. are considered one

of the most tolerant water-related pathogens to UV-C and PUV(Abshire and Dunton 1981; Chang et al. 1985; Hijnen et al.2006; Farrell et al. 2010). On the contrary, Staphylococcus spp.have often shown similar or greater sensitivity to UV technol-ogy than coliforms (Abshire and Dunton 1981; Chang et al.1985; Hijnen et al. 2006). However, the variability in UVresistance among species and strains is larger than the diver-gences among genera, which avoid general conclusions(Table 2). For instance, Abshire and Dunton (1981) reportedthat the resistance to UV-C of Enterococcus faecalis (ATCC10541) was 2.1 times higher than that of Staphylococcus au-reus (ATCC 6538). However, theDUV values of Pseudomonasspp. strains varied up to 4.0 times. Similarly, Farrell et al.(2010) observed that the difference in resistance to PUV be-tween E. faecium (NUIG J952) and S. aureus (NUIG 5624)was 2.0 times, whereas variations observed among P.aeruginosa strains were of the same order of magnitude.Higher divergences in UV-C resistance have been observedamong enteropathogenic E. coli strains, with varying DUV

values almost 13 times (Sommer et al. 2000). Due to the greatvariability in UV resistance among strains, the EnvironmentalProtection Agency's Scientific Advisory Panel has recom-mended establishing UV-C irradiation process criteria for waterdisinfection by testing five outbreak-related strains in a cocktailfor each pathogen (Oteiza et al. 2010).

Data on UV effectiveness against most significant food-borne pathogenic and spoilage bacteria are still limited.Listeria monocytogenes is considered one of the most UV-resistant foodborne pathogens in milk (Lu et al. 2011) andfruit juices (Guerrero-Beltrán and Barbosa-Cánovas 2006;Gabriel and Nakano 2009). On the other hand, Salmonellaspp. are one of the most sensitive genera (Gabriel and Nakano2009; Lu et al. 2011). For instance, Gabriel and Nakano(2009) reported a DUV value for L. monocytogenes (HCIPHAS-1) 2.1 times higher than for Salmonella typhimurium(ATCC 12529) in apple juice (Table 3). Nevertheless, a widervariability in UV resistance among strains of pathogenic bac-teria has been detected (Table 3), which may explain somediscrepancies in the literature when comparing the relativeresistance of species. A variation in DUV values of up to 5.5times among E. coli O157:H7 strains have been observed

Table 2 (continued)


Pseudomonas aeruginosa DIT 02B710 2.6c

NA data not available, LP low-pressure mercury vapor lamp, MP medium-pressure mercury vapor lamp, PX pulsed xenon lampaData estimated by Cairns (2006)b Values were estimated from different inactivation data from literature obtained using bench scale or continuous flow systems with one or more LP orMP lampscDUV values are expressed in number of light pulses (energy discharge 7.2 J)


(Gabriel 2012). More examples of UV resistance data forbacteria in liquid foods are provided in Table 3. Overall, moreinvestigation into UV resistance in different strains of relevantpathogens is needed in order to identify the most resistantmicroorganisms of public health significance.

Variations observed among microorganisms, species, andstrains have been attributed to different factors, including cellwall thickness; cell size; pigmentation; composition, size, and

conformation of the genetic material; and DNA repair efficien-cy. For example, the higher UV resistance of Gram-positiveversus Gram-negative bacteria has been related to the thickerpeptidoglycan cell wall of the former, which may hinder thepenetration of UV light photons within cells (Beauchamp andLacroix 2012).

Comparing the same genetic material content, larger sizedcells are more resistant to UV light because there is a higher

Table 3 UV doses required for 90 % reduction of the initial population(DUV values) of different types of microorganisms, species, and strainsreported in liquid foods. DUV values were expressed in fluence unit(millijoules per square centimeter), except where indicated. The type of

UV system (bench scale or continuous flow system) and UV source(number of lamps, type of lamps, and the total input power) used toobtain data are included


Saccharomyces cerevisiae BFE-39 Apple juice Bench scale 2× LP (15 W) 6.4a Gabriel (2012)Debaryomyces hansenii BFE-34 8.3a

Clavispora lusitaniae BFE-36 9.8a

Torulaspora delbrueckii BFE-37 9.4a

Pichia fermentans BFE-38 11.4a

Escherichia coli O157:H7 CR-3 2.8a

Escherichia coli O157:H7 HCPIH-96055 0.5a

Listeria monocytogenes HCIPH AS-1 Apple juice Bench scale 2× LP (15 W) 1.26a Gabriel and Nakano (2009)Listeria monocytogenes HCIPH M24-1 0.44a

Escherichia coli O157:H7 0.5a

Escherichia coli K12 0.6a

Salmonella typhimurium ATCC 12529 0.61a

Salmonella enteritidis HCIPH B11 0.27a

Saccharomyces cerevisiae Apple juice Thin film flow reactor 2× LP (25 W) 12.2 Guerrero-Beltrán andBarbosa-Cánovas (2006)Listeria innocua 4.9

Escherichia coli 2.9

Escherichia coli O157:H7 (303/00) Orange juice Bench scale 4× LP (30 W) 296 Oteiza et al. (2010)Escherichia coli O157:H7 (749/03) 169

Escherichia coli O157:H7 (547/03) 210

Escherichia coli O157:H7 (33/98) 168

Escherichia coli O157:H7 (EDL 933) 212

Saccharomyces cerevisiae DSM 70478 Apple juice Coiled tube (Vivatec) LP (8 W) 2.6b Müller et al. (2011)Alyciclobacillus acidoterrestris DSM

2498—vegetative cells1.3b

Escherichia coli DH5α 0.5b

Lactobacilllus plantarum BFG 5092 0.3b

Lactococcus lactis (cocktail) Whole milk Coiled tube LP (80 W) 5.1 Lu et al. (2011)Listeria monocytogenes (cocktail) 3.5

Escherichia coli (cocktail) 3.5

Salmonella typhimurium (cocktail) 3.4

Staphylococcus aureus (cocktail) 3.2

Pseudomonas aeruginosa (cocktail) 3.3

LP low-pressure mercury vapor lamp, MP medium-pressure mercury vapor lamp, PX pulsed xenon lamp

NA data not available, LP low-pressure mercury vapor lamp, MP medium-pressure mercury vapor lamp, PX pulsed xenon lampaData estimated by Cairns (2006)b Values were estimated from different inactivation data from literature obtained using bench scale or continuous flow systems with one or more LP orMP lampscDUV values are expressed in number of light pulses (energy discharge 7.2 J)


probability of photons being absorbed by other compoundsbefore affecting DNA (Oteiza et al. 2010; Gabriel 2012). Thisis one of the reasons molds and yeasts are more resistant thanbacteria (Tables 2 and 3). In addition, the lower pyrimidinecontent in the DNA composition of yeast may also contributeto its higher UV resistance (Fredericks et al. 2011).

Apart from DNA composition, the extent of UV damagehas been also correlated with genome size. Müller et al. (2011)attributed the larger UV-C resistance of S. cerevisiae comparedto Lactobacillus plantarum to higher cellular DNA content(Table 3). However, haploid yeasts appear to be more sensitiveto UV-C radiation than their diploid form (Rodrigues et al.2003), probably because target genes are present in severalcopies, decreasing the probability of all copies being hit.

DNA condensation also protects against UV damage.Deinococcus and Rubrobacter genera have strong UV resis-tance, which is partly attributable to their stronger genomecondensation compared with the uniform E. coli nucleoid(Blasius et al. 2008).

Differences in intrinsic repair efficiency may also contributeto the variations in UV survival among microorganisms. Cheiget al. (2012) attributed the greater UV-C and PUV sensitivity ofE. coli O157:H7 compared to L. monocytogenes to its smallerDNA repair activity, since their quantitative analysis of DNAdamage showed that the CPD formation in E. coliO157:H7wasslightly greater than in L. monocytogenes. Similarly, Suss et al.(2009) observed that the higher UV-C recovery ofP. aeruginosain comparison to a UV-sensitive strain of Enterococcus faeciumwas correlated with intensive repair in the former, whereas noDNA repair was detected in E. faecium.

The physical state (fluidity) of the cell membrane also canplay an important role in UV resistance because bacterialchromosomes interact with the cytoplasmic membrane. Someagents involved in DNA repair mechanisms (RecA and Uvrproteins) are attached to the membrane to perform their func-tions (Lin et al. 1997). It has been reported that excision repairis modulated by the fluidity of the cell membrane in UV-C-irradiated E. coli cells (Todo et al. 1983).

Finally, pigmentation is believed to protect cells by absorb-ing light at UV wavelengths and scavenging free radicalsgenerated by UV-A (Moeller et al. 2005). Thus, pyocyaninand pyoverdine pigments protect P. aeruginosa against UVlight (Burke et al. 1990; Farrell et al. 2010). However, thispoint should still be addressed because contradictory resultshave been found concerning the UV-protective role of pigmen-tation (Turtoi and Nicolau 2007; Khaneja et al. 2010).

Bacterial spores are well known for their great UV resis-tance properties. For example, spores of various Bacillus spe-cies are five to 50 times more resistant to UV radiation thanvegetative cells (Setlow 2001). This high resistance has beenascribed to several mechanisms: the relatively dehydrated stateof the spore core reducing the probability of pyrimidine dimer-ization, the effective action of SP lyase during germination, the

accumulation of a high percentage of dipicolinic acid in thedormant spore core (Slieman and Nicholson 2001), and thepresence of a thick spore protein coating (Riesenman andNicholson 2000). Finally, spores like Bacillus spp. are able toproduce a broad spectrum of pigments, especially carotenoids,during germination, which could provide some UV-A protec-tion (Moeller et al. 2005; Khaneja et al. 2010).

Extrinsic Factors

Among extrinsic factors influencing UV microbial resistance,the main ones are the growth phase, growing conditions,stressors prior to UV treatment, and recovery conditions afterprocessing.

Growth Phase The life cycle of bacteria includes periods ofgrowth at various rates interspersed with periods of nongrowth,depending on the availability of nutrients. UV resistance mayvary during these times. Several authors have reported thatstationary-grown cells in various bacterial species show en-hanced UV resistance compared to actively growing cells.For instance, Child et al. (2002) reported that the resistance ofmid-exponential phase cells of S. typhimurium grown in acomplete rich medium under UV-C (70 J/m2, UV Stratolinkerapparatus) in a buffer (pH 7) increased by more than 1.3 Log10cycles at the start of the stationary phase of growth. Most ofthese studies were performed in batch cultures where changesin other factors such as medium composition might havehidden the effect of the growth phase on UV sensitivity(Berney et al. 2006). Systematic studies of the effect of specificgrowth rate on E. coli's UV resistance have been conductedusing continuous cultures and show that the specific growthrate is strongly correlated with UV resistance (Berney et al.2006; Bucheli-Witschel et al. 2010). Bucheli-Witschel et al.(2010) reported that bacteria cultivated in glucose-MM atintermediate specific growth rates between 0.2 and 0.4 h−1

exhibited the lowest UV-C resistance and that it increasedabove and below these values. The difference in inactivationlevels between stationary growth cells of E. coli and cellsgrown in the chemostat culture at dilution rates of 0.02, 0.22,and 0.58 h−1 treated by UV-C (50 J/m2, bench scale device, 4×LP lamps—16 W) in the same growth medium were 0.5, 2.4,and 0.9 Log10 cycles, respectively. Considering that in batchcultures, the specific growth rate decreases constantly until thestationary growth phase is reached, UV-C resistance decreasedduring the mid-exponential phase and reached maximum resis-tance just before cells entered the stationary phase.

In Gram-negative bacteria, the increase in UV resistancealong with the progressive growth rate decrement has beenrelated to the induction of the general stress response sigmafactor RpoS (σ38) (Child et al. 2002; Berney et al. 2006;Bucheli-Witschel et al. 2010). Bucheli-Witschel et al. (2010)reported that RpoS is involved in the UV-C resistance of


growing E. coli cells at low rates (0.2–0.3 h−1) but not atmedium or high rates. This corresponds with the increase inRpoS expression when the growth rate decreases (Ihssen andEgli 2004). However, the sensibility of exponential phase cellscannot be explained simply by the low expression of RpoSfactor (Child et al. 2002; Bucheli-Witschel et al. 2010).Although there are some indications that relate the RpoSfactor to the action of the nucleotide excision repair system(Tyrrell et al. 1972), a direct relationship has not yet beenfound yet. A possible link between the RpoS factor and UV-Cresistance might be the induction of the expression of DNA-binding proteins (Dps). Dps bind to the chromosome, forminga stable nucleoprotein complex which may change DNAreactivity toward UV-C irradiation and increase the efficiencyand accuracy of repair mechanisms (Nair and Finkel 2004).

For high specific growth rates (>0.4 h−1), the decrease ofUV-C sensitivity in E. coli as the growth rate increases hasbeen related to expanding genome content. Bronk andWalbridge (1980) reported that the increase of UV-C resis-tance in exponential phase cells was due to the lengthening ofthe shoulder phase of survival curves when DNA content wasaugmented. According to the multiple target theory, this canbe attributed to the enhancement of a number of target mole-cules (Bronk and Walbridge 1980). In other words, the incre-ment of DNA content per cell may imply that a target gene ispresent in several copies, and therefore, it is less likely that allcopies will be hit and damaged (Bucheli-Witschel et al. 2010).Also, high activation of repair systems mediated by RecA dueto an increase in the number of replication forks could beinvolved (Dri and Moreau 1993).

InGram-positive bacteria, the effect of the growth phase hasnot been studied, with the exception of B. subtilis (Wassmannet al. 2011). The obtained results were the opposite of thegeneral assumption that exponentially growing cells are moresensitive to UV light than stationary-grown cells. The DUV

value of exponential phase cells of B. subtilis DE69 (2.4 h ofgrowth in nutrient broth) treated under a UV polychromaticlamp (200–400 nm, Sunlight Simulator SOL 2) in a buffer (pH7) was almost two times higher than that of stationary phasecells. This suggests that the expression of the stress sigmafactor B (SigB/σB) at the start of the stationary growth phaseis not involved in UV resistance. In fact, preliminary studiesfrom the same authors showed that the UV resistance ofstationary phase cells of a B. subtilis wild-type strain wasalmost identical to that of an isogenic σB mutant.

It is well known that during the stationary phase of bacterialgrowth, new genotypes appear through random spontaneousmutation (Dykhuizen 1999; Vulic and Kolter 2001). Genotypesthat can confer a competitive advantage in nutrient-poor con-ditions are selected, encouraging subpopulations of differentphenotypes (Vulic and Kolter 2001), which may have UVsensitivity that differs from the initial population. Child et al.(2002) observed that the inactivation of stationary-grown cells

of S. typhimurium subjected to prolonged incubation in batchcultures for 2 months showed a regular cycling of UV-Cresistance fluctuations, varying nearly 3 three Log10 cycles ata UV-C dose of 100 J/m2. Although RpoS is not involved inUV survivability at the late stationary growth phase (Child et al.2002; Bucheli-Witschel et al. 2010), it is indirectly relatedbecause it increases the mutation rate. RpoS stabilizes Pol IVin the stationary growth phase and also downregulates enzymesresponsible for mismatch repair (Foster 2007). Mutations mayresult in the appearance of UV-tolerant subpopulations that donot remain stable during prolonged incubation.

In summary, to draw concise conclusions when comparingthe UVresistance of different microorganisms or the influenceof related factors, it is necessary to specify the age of bacterialcultures. From a microbiological point of view, it would bevery interesting to thoroughly study the effects of growthphases on the UV resistance of different microbial groupsand to determine the intrinsic factors that could affect it.

Growing Conditions There is little data related to the growthfactors influencing microbial UV resistance; growing mediumcomposition and growth temperature are the primarily studiedparameters. Some authors have reported that cells grown inmedia with simple molecules arabinose or glycerol as carbonsources were more UV-C sensitive than those grown in glu-cose media (Rude and Alper 1972; Schenley et al. 1976).Schenley et al. (1976) demonstrated that the UV survivabilityof glucose-grown cells was higher than those grown in glyc-erol due to higher excision repair activity. These data suggestthat the metabolic activity of growth prior to UV irradiationdetermines the efficiency of DNA repair and thus UV resis-tance. However, the medium composition only varied the UV-C sensitivity of E. coli by 25 % (Schenley et al. 1976).

Regarding bacterial spores, it has been reported that spor-ulation conditions influence UV resistance. Bacillus spp.spores purified from soil exhibited two to three times higherresistance than B. subtilis spores grown in laboratory media(Nicholson and Law 1999). Therefore, giving precise indica-tions regarding the composition of growth and sporulationmedia is strongly recommended.

On the other hand, several have reported that indigenousfood microorganisms are more resistant to UV technology thanare pure cultures (Franz et al. 2009; Lu et al. 2011). Forinstance, UV-C treatment of raw milk at 10.7 mJ/cm2 appliedby a coiled tube reactor only reduced 2 Log10 cycles of thenative microorganisms, including Staphylococcus spp., Listeriaspp., Salmonella spp., and Pseudomonas spp. Meanwhile, thesame dose was able to inactivate more than 3 Log10 cycles ofeach pure culture inoculated in milk. To explain these differ-ences, the existence of subpopulations of native microorgan-isms with UV resistance genes turned on was proposed.

Variations in growth temperature induce changes in themembrane fatty acid composition of microorganisms to


maintain a degree of fluidity compatible with life (Casadeiet al. 2002). Therefore, since cytoplasmic membrane fluidityhas an essential role in DNA repair (Todo et al. 1983; Lin et al.1997), growth temperature could be a crucial factor in UVresistance. However, the available data indicate that stationaryphase cells of Cronobacter sakazakii (Arroyo et al. 2012b)and E. coli K12 (unpublished data) grown at 10 and 37 °Cshowed the same degree of UV-C resistance.

Stresses Prior to UV Treatments During food processing,bacteria may be exposed to stress conditions which can pro-duce DNA damage or interfere with replication, activating theSOS response (van der Veen and Abee 2011). As a conse-quence, cross-protection against subsequent UV treatmentscould occur. A great number of stressors encountered in somefood processing steps such as heat, acid, alkali, oxidative,osmotic, starvation, and ethanol have been shown to increaseUVresistance. In some cases, their effect is noticeable, increas-ing UV resistance up to 3.5 times after heat shocks (Pardasaniand Fitt 1989). This cross-protection is controlled by regulatorsthat induce the expression of multiple genes which aid micro-bial adaptation: alternative sigma factors (RpoS and RpoH inGram-negative bacteria and sigB in Gram-positive ones), smallmolecule effectors such as guanosine 5′-diphosphate 3′-di-phosphate (ppGpp), gene repressors such as LexA, and inor-ganic molecules such as polyphosphates (Foster 2007).Although the biology of stress responses in traditional foodhurdles has beenwidely studied, with regard to UV processing,there is little and contradictory information.

Exposure to oxidative agents such as hydrogen peroxideand sodium hypochlorite induces the intracellular formationof ROS, causing DNA damage and activating the SOS re-sponse (van der Veen and Abee 2011). For instance, previousexposure to oxidative stresses (2.5 mM H2O2 for 20 min)increased the UV-C survivability of E. coli cells treated at 12mJ/cm2 by around 0.5 Log10 cycles (Asad et al. 1994). Thisprotection involves the induction of some DNA repair mech-anisms dependent on RecA and UvrA proteins (Asad et al.1994). On the contrary, oxidative shock (H2O2 0.5 mM for 1h) sensitized C. sakazakii to UV-C (Arroyo et al. 2012a).Exposure to mildly acidic conditions (pH 5 for 3 h) providedcross-protection to L. monocytogenes against UV-C light,increasing survivability by 2.2 Log10 cycles in distilledwater after 20 min of irradiation (12.8–33.2 mJ/cm2)(McKinney et al. 2009). Similarly, the inactivation rate of L.monocytogenes to PUV light (7.2 μJ/cm2) decreased 25 %after an acid shock of pH 5.5 for 5 h (Bradley et al. 2012).However, no changes in the UV-C resistance of acid-stressedC. sakazakii (Arroyo et al. 2012a) or Lactococcus lactis(Hartke et al. 1995) cells were observed. Acid exposure resultsin DNA strand breaks, and the acid sensitivity increases inRecA-deficient strains, highlighting the role of the SOS sys-tem in acid resistance (van der Veen and Abee 2011).

Little information exists concerning the alkaline stress effecton UV sensitivity. Goodson and Rowbury (1990) observedthat the inactivation of E. coli and its RecA-deficient derivativegrown in nutrient broth of pH 9.0 was 53 % lower after 60 s ofUV-C irradiation than in those microorganisms grown at neu-tral pH. This could be explained by the inactivation of LexArepressor under alkaline conditions (Little 1991). In contrast,UV-C resistance of C. sakazakii was significantly reduced (20%) after an alkaline shock at pH 9 for 1 h (Arroyo et al. 2012a).Exposure to high salt concentrations results in the modulationof the structural conformation of RecA to an activated state(Dicapua et al. 1990), which could turn on the SOS responseand boost microbial UV resistance. However, osmotic shocksdid not change the sensitivity of L.monocytogenes to PUV (7.5% NaCl for 1 h) (Bradley et al. 2012) or ofC. sakazakii to UV-C (5 % NaCl for 1 h) (Arroyo et al. 2012a). Alternatively, pre-irradiation of E. coli cells with monochromatic visible light atwavelengths of 446, 466, 495, and 685 nm for 30 min in-creased UV-C cell survivability (12mJ/cm2) by almost 1 Log10cycle (Lage et al. 2000).

Heat shock (HS) is one of the most studied stressors thatcould affect microbial survivability to UV light. For instance,HS at 42 °C for 45 min increased the UV-C survivability (35–60 J/m2) of E. coli up to 3.5 times (Pardasani and Fitt 1989).Similarly, Lage et al. (2000) observed a decrease of 1.9 Log10cycles in the UV-C inactivation (120 J/m2) ofE. coliwhen cellswere pre-irradiated with polychromatic infrared radiation (60min). This was attributed to the localized temperature increaseswithin the cellular environment. The UV resistance incrementsafter HS are related to the stimulation of excision repair via theSOS response (Pardasani and Fitt 1989). However, other au-thors noticed no PUVor UV-C survival improvement (Arroyoet al. 2012a; Bradley et al. 2012). Moreover, L.monocytogenes(McKinney et al. 2009) and Vibrio cholerae (Jeevan andGhosh 1995) became more vulnerable. When bacteria aresubjected to a rapid increase in external temperature, theyrespond by expressing heat shock proteins (HSPs) under thetranscriptional control of the alternative sigma factor RpoH(σ32), increasing their subsequent thermotolerance. HSP re-sponse overlaps with SOS response by protecting heat-labileproteins important for DNA repair from degradation (Zou et al.1998; Layton and Foster 2005).

UV survival improvement after starvation stresses has alsobeen widely reported. For instance, the UV-C resistance (18mJ/cm2) of E. coli after 24 h of deprivation of amino acids orthiamine and glucose increased by about 0.75 and 1.75 Log10cycles, respectively, in comparison with cells grown in acomplete medium (Fitt and Sharma 1991). In this case, con-trary to what was expected, the RpoS response under nutrientlimitation conditions hardly contributed to this phenomenon.The implications of the RpoH response and ppGpp in tempo-rarily halting DNA replication have been suggested (Sedliakova1998; Foster 2007).


Finally, homologous protection after the application oflow UV doses has been observed, a phenomenon calledphotoprotection. Pre-exposure to a near-UV lamp (300–400nm, 8×104 J/m2) causes DNA lesions that trigger the SOSresponse and growth delay as part of the SOS function(Bhatnagar 1992). On the other hand, low UV doses (50–100J/m2) induce HSP synthesis (DnaK and GroEL proteins) in E.coli, contributing to UV resistance improvement (Krueger andWalker 1984). Similarly, pretreating P. aeruginosa with low-energy doses of PUV (0.2 J/m2) increased the survivability offurther high intensity PUV treatments (1 J/cm2) by 3 Log10cycles (Massier et al. 2012). Overall, as the development ofcross-protection against UV light during food processing couldhave important consequences in terms of food safety, it isnecessary to obtain deeper knowledge concerning the stressfulconditions that improve microbial UV resistance and under-stand the causes of existing discrepancies.

Recovery Conditions After Processing Environmental condi-tions after UV treatment can determine microbial resistance toa great extent. Bacteria survivability after UV irradiation ishigher when cells are plated on minimal growth medium ratherthan on rich growth medium (Ganesan and Smith 1968).Similarly, cell recovery in a minimal liquid medium in thedark, that is, so-called liquid holding recovery (LHR), is betterthan when the cells are directly plated in complete agar. Thisphenomenon has been attributed to NER action before plating(Ganesan and Smith 1968).

Based on Ganesan and Smith's first hypothesis (1968), thenumber of dimers that can be removed by NER depends onthe number of enzymes present in the irradiated cells and thetime available for NER before replication ensues. In completegrowth media, the maximum amount of time available forNER would be one generation time. If a cell is not able toremove all dimers, then replicational repair pathways will thenbe activated, including error-prone mechanisms. Thus, cellswill have more probability of DNA repair failure, resulting innonviable cells. This conjecture was set aside when moleculartechniques appeared, allowing researchers to thoroughly studythe DNA repair pathways. However, from a practical point ofview, it is critical to know which repair mechanisms can occurin different recovery media, such as food matrices, and espe-cially to understand their impact on bacteria survivability.Therefore, these investigations should be reconsidered.

Currently, LHR is important due to the increase in thenumber of UV treatment facilities for drinking water andwastewater treatment throughout the world. Thus, the recov-ery of pathogens in dark conditions after applying UV dosesnow considered safe might make treatment ineffective. Theextension of LHR depends on the intrinsic abilities of eachmicroorganism, which vary among species and even strains.For instance, LHR plays no role or a minimal one in E. coli

recovery (Sommer et al. 2000), with the exception of strainO50:H7. The UV-C dose required to achieve 6 Log10 reduc-tions of E. coli O50:H7 in clear buffer increased 1.7-foldconsidering LHR (48 h at 22 °C). Pathogenic species in fishsuch as Aeromonas spp. and Vibrio spp. are able to carry outextensive LHR. Aeromonas salmonicida subsp. salmonicidashowed a dark recovery (48 h at 22 °C) of almost 2 Log10cycles after applying a UV-C dose of 6 mJ/cm2 (3 Log10cycles of inactivation) (Liltved and Landfald 1996). LHR istemperature dependent, occurring to a greater extent at 37 °Cthan at lower temperatures (4 °C) (Liltved and Landfald1996). LHR has been well described when microorganismsare treated in water. However, it has hardly been investigatedin treated foods, which is an essential point in applying UVtechnology to food hygienization.

In addition to dark repair, photoreactivation can be anefficient DNA repair mechanism and much more importantthan dark repair (Liltved and Landfald 1996; Sommer et al.2000). Photoreactivation is crucial for water disinfection whenwater is exposed to visible light after UV treatment. However,it has been poorly considered whenmicroorganisms are treatedin food.

The photorepair capability of microorganisms differs greatlyfor different species and strains (Tosa and Hirata 1999; Hassenet al. 2000; Sommer et al. 2000). This ability depends onwhether the enzyme photolyase gene is present, and in bacteria,its degree of expression is distributed in an unpredictablemanner. Hassen et al. (2000) reported that UV-C irradiated cells(0.3 J/cm2) of Serratia marcescens, Acinetobacter baumanni,and Enterobacter aerogenes were able to repair more than 5Log10 cycles of viable cells after visible light illumination(24 h), whereas P. aeruginosa, which showed the highest UV-C resistance, was not able to photoreactivate. The photorepairability of different strains of enteropathogenic E. coli strains hasbeen widely studied, with great variability observed. For in-stance, Tosa and Hirata (1999) observed that the maximumUVdose required for 90 % UV-C inactivation of E. coli O26 withphotoreactivation (0.023mW/cm2 at 360 nm for 4.6 h) was 2.2-fold times higher than that without photoreactivation.

Besides the microbially specific response, photoreactivationdepends on the delivered UV dose, the UV irradiation source,and the visible light illumination conditions. The extent ofphotoreactivation is often inversely related to the applied UV-C dose (Lindenauer and Darby 1994). Thus, applying suffi-ciently high UV doses, photoreactivation can be drasticallyreduced. The use ofMP instead of LP lamps has been proposedas another strategy to control photoreactivation. Some studieshave demonstrated that MP sources resulted in lower photore-activation levels of E. coli than LP lamps had for the samegermicidal UV dose (up to 10 mJ/cm2) (Oguma et al. 2002;Zimmer and Slawson 2002). Halmich and Gehr (2010) attrib-uted this phenomenon to simultaneous exposure to UV-C light


and visible light (400–800 nm). On the contrary, other authorshave found similar degrees of E. coli photoreactivation for bothMP and LP lamps after treatments of 40 mJ/cm2 (Guo et al.2009). Furthermore, photoreactivation has been observed aftermicrobial inactivation by the broad spectrum of PX lamps(Maclean et al. 2008). Therefore, processing conditions thatavoid photoreactivation should be better addressed to establishadequate operational parameters.

Regarding post-illumination conditions, photoreactivationtakes place above a certain visible light intensity threshold.According to Halmich and Gehr (2010), at least 440 l × (0.065mW/cm2) of visible light is needed to initiate E. coli photoreac-tivation. Once this threshold is surpassed, the photorepair rateincreases with the visible light intensity (Tosa and Hirata 1999).However, the maximum survival obtained after photoreactiva-tion is independent of the intensity level (Lindenauer and Darby1994; Kashimada et al. 1996), indicating that it is the illumina-tion time which determines the final recovery value. This isbecause the limiting factor in the photorepair mechanism is thefrequency of photolyase attachment to the CPDs (Gehr andWright 1998). The higher the intensity of visible light, the higherthe probability of photons reaching the photolyase enzyme.Moreover, the emission spectrum of visible light sources alsoaffects the photorepair rate (Bohrerova and Linden 2007;Maclean et al. 2008). However, by comparing photorepair ratesbased on photorepair fluence calculations (weighted at 300–500nm), Bohrerova and Linden (2007) demonstrated that these rateswere independent of the lamp type, but larger differences couldbe found using time-based comparisons. In addition, the delay invisible light exposure after UV treatment is also important(Lindenauer and Darby 1994; Hallmich and Gehr 2010).Delaying exposure to visible light for 3 h suppressed photore-activation after relatively low UV-C doses (10 and 20 mJ/cm2)(Hallmich andGehr 2010). Overall, to compare the repair abilityof different microorganisms, post-illumination conditions afterUV treatment should be carefully optimized to attain the max-imum expression of photorepair ability.

Environmental factors such as temperature, pH, and ionicstrength influence the rate of the photoreactivation process.Temperature has little effect on bacterial photoreactivation,but it occurs faster at higher temperatures (37 °C) than atlower ones (4 °C) (Liltved and Landfald 1996). The effect ofsalinity seems to be greater than that of temperature. Thebinding affinity of the CPD photolyase decreases under lowionic strength conditions, and consequently, photoreactivationdecreases (Chan and Killick 1995).

Although photoreactivation is important in water disinfec-tion, in food processing, photorepair could not play such animportant role because foods are normally stored andtransported under dark conditions in containers. However, itshould be considered when UV-processed foods are likely tobe exposed to visible light.

Product Parameters

Other characteristics of the treatment media may change thebactericidal efficacy ofmost food preservation technologies. Ingeneral, the pH and water activity of the treatment media arethe most investigated factors. Several authors have reportedthat the lethality of UV treatments is independent of pH(Quintero-Ramos et al. 2004; Gayán et al. 2011). This islogical because the key target of UV light is DNA, so it wouldaffect cell envelopes minimally. However, the effect of wateractivity (aw) has barely been investigated. It has been demon-strated that water activity does not alter the UV resistance of E.coli or S. typhimurium in liquidmediumbetween 0.96 and 0.99(Gayán et al. 2011, 2012a). Similarly, the water activity of foodpowder (black pepper and wheat flour) decontaminated byPUV did not affect the inactivation of S. cerevisiae between0.3 and 0.6 (Fine and Gervais 2004). From a practical point ofview, the absence of effects from pH or aw on bacterial UVresistance is of extreme importance because it facilitates theapplication of this technology to any food product indepen-dently of these factors.

The most influential product characteristics in terms of thelethal efficacy of UV technologies are the optical properties,mainly the UV absorbance and the turbidity of the medium.Color components, soluble compounds, and suspended solidscan absorb, reflect, and scatter the incidental light, reducing thenumber of photons available for killing microorganisms(Koutchma 2009). This can be observed by comparing UVresistance data from Tables 2 and 3, whichwere obtained usingclear media and food, respectively. The absorption coefficient(α) is considered a good indicator of UV penetration depth inclear liquids, and several authors have reported a good rela-tionship between α and UV lethal efficacy (Koutchma 2009;Gayán et al. 2011). For instance, Gayán et al. (2011) demon-strated a linear relationship between α and the Log10 of theinactivation rate of E. coli: The inactivation rate decreased 10times by increasing the absorption coefficient by 15.9 cm−1.The negative impact of suspended solids on UV decontamina-tion has been also reported, but its direct effect on UV lethalityis not so clear (Koutchma 2009). In any case, although theoptical properties of foods can affect microbial UV resistanceto a great extent, their effects on UV light's lethal effectivenessare due to light intensity attenuation rather than to directbiological effects.

Combination Processes

The limited microbial lethality of UV light in media with ahigh absorption coefficient and turbidity, such as most liquidfoods, promotes the development of combined processes. UVtechnology can be combined with conventional and other


nonthermal processes to enhance the lethal effects of UV lighton microorganisms. The lethality of combined treatments canresult from an additive or synergistic effect. Synergistic lethaleffects are preferable when designing combined processesbecause they allow for obtaining a specific level of inactiva-tion, reducing the energy input and treatment intensity. Thissynergy may arise when applied inactivation strategies act onthe same vital target, increasing the chances of irreversibledamage, or when they act on different sites, causing sublethaldamage in several areas of the cells, enhancing their overalllethal potential (Ross et al. 2003). The intelligent design ofcombined treatments requires knowledge of the mechanismsof inactivation for each technology to be applied. A synergisticeffect has been observed in combining UV light with mildheat, chemical agents, and ionizing radiation (Gentner andWerner 1978; Gayán et al. 2011). However, the combinationof UV light with other nonthermal technologies such as pulsedelectrical fields or ultrasounds for fruit juice hygienization hasresulted in additive inactivation effects (Gachovska et al. 2008;Char et al. 2010).

When combining UV light with cooling or mild heating,first of all, it is important to distinguish the changes in UVmicrobial survivability due to biological effects from thosecaused by variations in the efficiency of UV sources at differ-ent temperatures. The optimum temperature of 254 nm radia-tion produced by LP lamps is 40 °C, and then it drops off withincreased temperature (Koutchma 2009).

Besides the physical influence of temperature on UV le-thality, reducing treatment temperature has been associatedwith lower UV inactivation. For instance, Oteiza et al. (2010)compared the UV-C lethal efficacy of E. coli O135:H7 strain33/98 in orange juice at 4 and 20 °C. Results demonstratedthat the DUV value obtained at 4 °C (168 mJ/cm2) was greaterthan at 20 °C (191 mJ/cm2). The higher UV microbial resis-tance at lower temperatures could be explained by the minornumber of pyrimidine dimers formed in vivo and by changesin the interactions between DNA and DNA-binding proteins,which help to maintain the correct functional DNA structure(Matallana-Surget et al. 2010).

Above 40 °C, the synergistic lethal improvement of UV-Clight when treatment temperature is increased by up to 50–60°C has been reported in laboratory media and food matrices(Gayán et al. 2011, 2012a, b). For instance, UV-C inactivation(27.10 J/mL) of E. coli in orange juice increased synergisti-cally from 0.6 Log10 cycles at 25 °C to more than 6 Log10cycles at 55 °C. Heat inactivation at this temperature and forthe same treatment time was negligible (Gayán et al. 2012b).

The mechanism of the synergistic effect of the combinedUV light and mild heating treatments (UV-H treatments) re-mains unclear. Our preliminary results demonstrated that thesynergistic lethal effect of UV-H treatment on E. coli resultedfrom the inhibition of DNA excision repair due to membranefluidification caused by simultaneous heating (Gayán et al.

2013). In addition, we reported that the number of envelope-injured cells was higher after the UV-H treatment than afterheating alone, indicating that the sensitization of cell envelopesor inability of cells to repair these structures also occurs (Gayánet al. 2012a).

UV light also exhibits synergistic interactions in combinationwith various chemical agents. In practice, such combinationshave been introduced in water disinfection as advanced oxida-tion processes (AOPs). AOPs are based on the utilization ofsecondary oxidants such as hydroxyl radicals, released by UVphotolysis decomposition of a chemical disinfectant to improvethe UV germicidal effect and the oxidation of organic com-pounds. Paracetic acid (PAA, 0.5–1.5 mg/L) in combinationwith UV-C light (8–10 mJ/cm2) can improve lethal efficacy onpathogen vegetative bacteria synergistically (Koivunen andHeinonen-Tanski 2005). Koivunen and Heinonen-Tanski(2005) observed 2.2 Log10 cycles of synergistic inactivation ofE. coli by combining 3 mg/L of PAAwith a UV-C dose of 10 J/cm2. Alternatively, the combination of ozone (3.4 mg min/L)and UV-C light (14 mJ/cm2) showed a synergistic lethal effect(2.5 Log10 cycles of inactivation) on B. subtilis spore inactiva-tion (Jung et al. 2008). Other chemical agents that can enhanceUV-C lethal efficacy include TiO2 (Wu et al. 2011) and ethanol(Ha and Ha 2010). In general, a simultaneous combination hasgreater lethal synergism than sequential treatments, probablydue to increased radical formation, and this synergy increaseswhen either the UV level or the chemical dose is raised(Koivunen and Heinonen-Tanski 2005; Jung et al. 2008).Overall, the combination of UV light with oxidant agents allowsfor reduced disinfectant doses or treatment times required forwater disinfection, thereby decreasing processing costs.

Combining UV-oxidant agents for food hygienization hasbeen less studied, but this has been proven to decontaminateeggshells (Rodriguez-Romo and Yousef 2005), seafood (Ramoset al. 2012), and fruit and vegetable surfaces (Fonseca andRushing 2006) without affecting food quality. For instance,Rodriguez-Romo and Yousef (2005) demonstrated that thecombination of UV-C light (90–150mJ/cm2) followed by ozone(12–14 % for 1 min) resulted in synergistic inactivation ofSalmonella enteritidis by 4.6 Log10 units or more.

Although interesting results have been found by combiningUV light with physical and chemical agents, more research isneeded to establish effective combined processes for foodhygienization. For this purpose, it is essential to know theaction mechanism for each hurdle as well as the environmen-tal, process, and microbial factors that influence microbialinactivation and cell damage.

Conclusion

Nowadays, rising interest in the use of UV-based technologiesfor food processing has prompted researchers to study the


lethal effect of these technologies on foodborne microbes. UVlight's ability to inactivate a wide range of microorganisms hasbeen extensively reported. However, little information on theUV resistance of significant foodborne and spoilage bacteria isavailable. Therefore, extending existing data is necessary toidentify key pathogens of concern in order to design UVinactivation processes. The high variability in UV resistanceobserved among strains of pathogenic species, which is evenbroader than the interspecies and intergenera variability, shouldbe considered. The age of growth and the growth conditionsfor bacteria may exert important effects on UV sensitivity, andtherefore, they should be standardized so microbial resistancecan be properly compared. In addition, knowledge ofmicrobialfactors that contribute to the high resistance of UV-tolerantmicroorganisms, such as chromosome condensation or DNArepair efficiency, could be useful in creating strategies thatimprove UV lethal efficacy, for instance, irradiating cells underadverse conditions to impair DNA repair.

From a practical point of view, UV sensitivity is an intrinsiccharacteristic of each microorganism. However, there are otherextrinsic microbial factors, processing conditions, and foodcharacteristics which also affect the UV resistance to someextent. Knowing the influence of some of these factors onUV survivability is crucial for efficient microbial control.Discrepancies in the effect of sublethal stressors prior to UVtreatment must be evaluated in order to prevent adverse situa-tions that might endanger food safety. At the same time, recov-ery conditions strongly determine microorganisms' capacity torepair UV-induced damage and survive. Of course, exposure tovisible light after UV processing must be completely avoided.Furthermore, the ability of microorganisms in foods to performwhat is known as “dark repair” should be evaluated.

Finally, the possibility of designing combined treatmentsbased on UV light and other physical or chemical agents hasbeen addressed. Deeper knowledge of the mechanisms behindUV inactivation and the functioning of damage repair pathwayscan help researchers to design intelligent treatment combina-tions. Thus, the simultaneous combination of UV light withmild heating, oxidant agents, or cell membrane-fluidificantcompounds may result in successful inactivation treatments.

Acknowledgments This study has been carried out with financial supportfrom the Ministerio de Ciencia e Innovación de España, EU-FEDER(CIT020000-2009-40) and the Departamento de Ciencia, Tecnología yUniversidad del Gobierno de Aragón. E. G. gratefully acknowledges thefinancial support for her doctoral studies from theMinisterio de Educación yCiencia de España.

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