Biotechnology Advances 28 (2010) 308–324
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Biotechnology Advances
j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / b i o t e c h a d v
Research Review Paper
Reactor design for minimizing product inhibition during enzymatic lignocellulose
hydrolysis: I. Significance and mechanism of cellobiose and glucose inhibition on
cellulolytic enzymes
Pavle Andrić, Anne S. Meyer ⁎, Peter A. Jensen, Kim Dam-Johansen
Department of Chemical and Biochemical Engineering, Technical University of Denmark, DK-2800 Kgs. Lyngby, Denmark
a r t i c l e
i n f o
Article history:
Received 15 October 2009
Received in revised form 28 December 2009
Accepted 7 January 2010
Available online 18 Janaury 2010
Keywords:
Cellulases
Lignocellulose
Product inhibition
Enzyme kinetics
Product removal
Reactor design
a b s t r a c t
Achievement of efficient enzymatic degradation of cellulose to glucose is one of the main prerequisites and
one of the main challenges in the biological conversion of lignocellulosic biomass to liquid fuels and other
valuable products. The specific inhibitory interferences by cellobiose and glucose on enzyme-catalyzed
cellulose hydrolysis reactions impose significant limitations on the efficiency of lignocellulose conversion —
especially at high-biomass dry matter conditions. To provide the base for selecting the optimal reactor
conditions, this paper reviews the reaction kinetics, mechanisms, and significance of this product inhibition,
notably the cellobiose and glucose inhibition, on enzymatic cellulose hydrolysis. Particular emphasis is put
on the distinct complexity of cellulose as a substrate, the multi-enzymatic nature of the cellulolytic
degradation, and the particular features of cellulase inhibition mechanisms and kinetics. The data show that
new strategies that place the bioreactor design at the center stage are required to alleviate the product
inhibition and in turn to enhance the efficiency of enzymatic cellulose hydrolysis. Accomplishment of the
enzymatic hydrolysis at medium substrate concentration in separate hydrolysis reactors that allow
continuous glucose removal is proposed to be the way forward for obtaining feasible enzymatic degradation
in lignocellulose processing.
© 2010 Elsevier Inc. All rights reserved.
Contents
1.
2.
3.
4.
5.
6.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Reactor goals: Enzymatic cellulose hydrolysis . . . . . . . . . . . . . . . . . . .
Enzymatic cellulose hydrolysis reactions . . . . . . . . . . . . . . . . . . . . .
3.1.
The Trichoderma reesei cellulase system . . . . . . . . . . . . . . . . . .
3.2.
Inhibition of cellulolytic enzymes by cellobiose and glucose . . . . . . . . .
Inhibition schemes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4.1.
Common inhibition kinetics . . . . . . . . . . . . . . . . . . . . . . . .
4.2.
Product inhibition . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4.3.
Diminishing the (product) inhibition effect . . . . . . . . . . . . . . . . .
Cellulolytic enzyme inhibition studies and results . . . . . . . . . . . . . . . . .
5.1.
Glucose inhibition of cellulolytic enzymes . . . . . . . . . . . . . . . . .
5.2.
Glucose inhibition of β-glucosidases . . . . . . . . . . . . . . . . . . . .
5.3.
Inhibition by cellobiose . . . . . . . . . . . . . . . . . . . . . . . . . .
5.4.
Cellulase inhibition terminology controversies . . . . . . . . . . . . . . .
5.5.
Mathematical curiosity of non-competitive modeling of cellulolytic inhibition
Experimental problems of enzymatic cellulose hydrolysis in relation to understanding
6.1.
Product inhibition study strategy . . . . . . . . . . . . . . . . . . . . . .
6.2.
The cellulosic substrate . . . . . . . . . . . . . . . . . . . . . . . . . .
6.3.
The cellulolytic enzymes . . . . . . . . . . . . . . . . . . . . . . . . . .
6.4.
Enzyme-to-substrate ratio (E/S) . . . . . . . . . . . . . . . . . . . . . .
⁎ Corresponding author. Tel.: + 45 4525 2800.
E-mail address: [email protected] (A.S. Meyer).
0734-9750/$ – see front matter © 2010 Elsevier Inc. All rights reserved.
doi:10.1016/j.biotechadv.2010.01.003
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6.5.
Substrate inhibition and transglycosylation . . .
Unique features of cellulase action and inhibition . . .
7.1.
Cellulase binding domain (CBD) and inhibition .
7.2.
Cellulase catalytic domain (CD) and inhibition .
7.3.
Unique parameters affecting cellulase inhibition
7.4.
Consequences for reactor design . . . . . . . .
8.
Minimizing cellulase inhibition: Conclusions . . . . .
References . . . . . . . . . . . . . . . . . . . . . . . .
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1. Introduction
The effective generation of a fermentable hydrolysate from cellulose
is one of the main requirements for cost-competitive production of
biofuels and other sustainable biobased products from lignocellulosic
feedstocks (Himmel et al., 2007; Lynd et al., 2008). Intense research
efforts are currently directed towards improving the hydrolytic
degradation of lignocellulosic biomass. These efforts include improvements of both the biomass pre-treatment technologies and the
cellulolytic enzymes that catalyze the conversion of the (ligno)cellulose
to glucose and pentoses (Merino and Cherry, 2007; Mosier et al., 2005).
Significant progress has already been made with respect to
selecting and developing better enzymes with improved stability,
higher specificity, and faster action on solid lignocellulose substrates
(Merino and Cherry, 2007; Zhang et al., 2006), but improvement of the
enzymatic hydrolysis is still considered one of the main research
challenges for achieving cost-effective lignocellulose-to-ethanol processing (Hahn-Hägerdal et al., 2006; Himmel et al., 2007).
Simultaneous saccharification and fermentation (SSF), with or
without separate fermentation of pentose monosaccharides, is at
present considered a main current technology scenario for biomass
conversion (Hahn-Hägerdal et al., 2006; Lynd et al., 2008). Although the
product inhibition of the enzymes is a rationale for SSF, the efficiency of
this technology is restricted by the (often overlooked) inhibition of
ethanol on the cellulolytic enzymes (Bezerra and Dias, 2005; Wu and
Lee, 1997). In addition, SSF inherently compromise the rate of the
enzymatic hydrolysis because of the relatively low temperature
requirement, i.e. 30–32 °C, for the fermentation. Hence, a certain degree
of separate enzymatic hydrolysis of cellulosic biomass is likely to be the
most feasible approach in future large-scale cellulose-to-ethanol or
other lignocellulosic biomass upgrading processes.
Unfortunately, the currently employed cellulolytic enzyme systems,
that include the widely studied Trichoderma reesei enzymes, are
significantly inhibited by the hydrolysis products cellobiose and glucose.
This inhibition retards the overall conversion rate of lignocellulose-toglucose (Gan et al., 2002; Katz and Reese, 1968). The significance of this
product inhibition is particularly prominent during processing at high
substrate loadings, not only because the enzymatic reactions may occur
faster at high substrate levels — given the same enzyme concentration,
but notably because the glucose concentration – all other things being
equal – will increase to a higher final level in the volume of liquid at high
dry matter loads than at low dry matter loads (Kristensen et al., 2008a;
Rosgaard et al., 2007a). This problem currently limits the extents of
conversion and the glucose yields achievable in batch processing of
lignocellulose (Rosgaard et al., 2007b; Tengborg et al., 2001), and
provides a large incentive to optimize the reactor designs to alleviate the
product inhibition and maximize the enzymatic conversion. The
significance of the conversion efficiency and hence the reactor design
is particularly important in large-scale processing of cellulosic biomass,
where the costs of the conversion processing are a significant parameter
(Lynd et al., 2008; Wyman, 2008).
This paper reviews the significance of the product inhibition on
cellulolytic enzyme hydrolysis notably by cellobiose and glucose on
fungal cellulases. The main purpose of the treatise is to understand the
significance of this inhibition in relation to designing reactors and
reaction systems that minimize or alleviate it. A second purpose is to
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examine the mechanistic features of inhibition in relation to the
enzymatic hydrolysis reactions. Alleviation of the product inhibition
may be accomplished via glucose removal to allow maximal enzymatic
conversion rates, optimal use of the enzymatic catalysts, high volumetric
productivity, and high glucose yields. The present review is tightly
connected to another report which examines the bioreactor designs and
reaction regimes – mainly membrane bioreactor strategies – that may be
suitable for product removal of glucose during cellulolytic hydrolysis
reactions (Andrić et al., submitted for publication). Our overall aim is to
highlight the immense potential – as well as the challenges – that lie
ahead in the development of reactor and process designs that reduce
product inhibition of cellulases.
2. Reactor goals: Enzymatic cellulose hydrolysis
A major requirement in cost-efficient lignocellulosic biomass
processing is to employ reactor systems that will ensure, or even
promote, maximal conversion of the cellulose with minimal enzyme
dosage, as the enzymes are still one of the single most important cost
terms in lignocellulose-to-ethanol processing (Hahn-Hägerdal et al.,
2006; Lynd et al., 2008). In practical terms, the effective use of the
catalyst, i.e. the yield of glucose obtained per amount of enzyme(s), in
lab-scale experiments often referred to as gglucose/gcellulase, and the
efficient bioconversion of the substrate itself, designated as gglucose/
gcellulose, are considered the principal parameters of importance for the
design and operation of appropriate bioreactor systems for lignocellulose conversion. The maximization of the product concentration, i.e.
the amount of glucose obtained per liquid volume, (kgglucose/m3), is
also considered a crucial parameter. In addition, optimization of the
volumetric productivity, in this case the rate of glucose formation per
reactor volume, kgglucose/(m3reactor volume h), is of profound significance
for achieving economically viable and sustainable cellulosic biomass
processing, especially on the large-scale and in a continuous mode.
Compared to other enzyme-catalyzed reactions the volumetric
productivities of cellulose conversion reactors are limited by the
inherently slow rate of the enzyme-catalyzed degradation of (ligno)
cellulose to glucose (Gan et al., 2002). For non-complexed cellulase
systems, which are in focus in this review, this slow rate is mainly due
to the insoluble and partly crystalline nature of the cellulose substrate
(Himmel et al., 2007; Zhang and Lynd, 2004). Nevertheless, within
these constraints, a reasonably high volumetric productivity is still a
prerequisite for ensuring reasonable equipment sizes in e.g. the largescale, continuous production of bioethanol. Hence, as we have outlined
in detail in a separate report (Andrić et al., submitted for publication),
the required size of the bioreactor equipment will increase for a certain
production goal if the volumetric productivity is low. A too low rate of
glucose formation per reactor volume may even prevent feasible
processing as it can require unrealistic reactor dimensions, (Andrić et
al., submitted for publication). The cellulolytic degradation accomplished by the complexed cellulase systems, i.e. the cellulosomes, that
have mainly been studied from different clostridia including Clostridium thermocellum, Clostridium cellulolyticum, Clostridium cellulovorans, and Clostridium josui will not be discussed here as it has been
reviewed elsewhere (Schwarz, 2001).
The quest for obtaining high glucose levels in lignocellulose-tobioethanol processing is mainly a result of the requirements related to
310
P. Andrić et al. / Biotechnology Advances 28 (2010) 308–324
the ethanol distillation step (Fig. 1), which demands relatively high
ethanol concentrations to be energy-efficient; in practice – as a
minimum – >4–5 wt.% (kg ethanol per 100 kg solution); which is
equivalent to ≈5–6% (v/v) (Alzate and Toro, 2006; Galbe et al., 2007;
Hahn-Hägerdal et al., 2006). Clearly, processing of high dry matter
contents, hence high cellulose levels, will also drive up the product
concentration (Fig. 1). This logic has directed some research into
attempting the enzymatic biomass conversion at high-solids and dry
matter levels (Jørgensen et al., 2007; Tolan, 2002). At high solid loadings,
the viscosity of the reaction mixture will be very high, and other factors
than product inhibition, notably mixing and mass transfer limitations,
and presumably increased inhibition by intermediates, also come into
play. Various fed-batch strategies for supplying the substrate to avoid
excessive viscosities and unproductive enzyme binding to the substrate
have been attempted (Rosgaard et al., 2007a; Rudolf et al., 2005).
Although high extents of lignocellulosic substrate conversion have been
achieved, the volumetric productivities have been low. When basing the
calculations on a product concentration of e.g. 86 g/L obtained in a batch
hydrolysis process, run for 4 days (96 h) at 40% dry matter of pretreated wheat straw (Jørgensen et al., 2007), and assuming that the
reaction mixture took up 2/3 of the reactor and that the reaction volume
reduced to 50% of the original reaction volume at the end of the reaction,
the volumetric productivity was <0.3 kgglucose/(m3reactor volume h).
3. Enzymatic cellulose hydrolysis reactions
The enzymatic hydrolysis of cellulose is a multi-step reaction that
takes place in a heterogeneous system, in which insoluble cellulose is
initially broken down at the solid–liquid interface via the synergistic
action of endoglucanases (EG) (EC 3.2.1.4) and exo-glucanases/
cellobiohydrolases (CBH) (EC 3.2.1.91) (Kubicek, 1992; Zhang and
Lynd, 2004). This initial degradation is accompanied by further liquidphase hydrolysis of soluble intermediate products, i.e. short cellulooligosaccharides and cellobiose, that are catalytically cleaved to
produce glucose by the action of β-glucosidase (BG) (EC 3.2.1.21)
(Fig. 2, paths 1 and 2) (Kubicek, 1992; Zhang and Lynd, 2004).
In practical lignocellulose processing, the cellulose being exposed to
enzymatic hydrolysis has usually undergone some kind of physicochemical pre-treatment that results in the cellulose consisting of both
Fig. 1. Theoretical (max. achievable) glucose concentration, g⋯L− 1 aqueous phase, after a
batch hydrolysis step and ethanol concentration % (volume/volume) after the batch
fermentation (SHF), as a function of pre-treated lignocellulose dry matter content (DM%).
The graphs were calculated from mass balances, reactions stoichiometry, an assumption of
100% yield in the hydrolysis and fermentation steps, and 50% (w/w) cellulose fraction in
the pre-treated material. The amount of water consumed by the reaction has been
accounted for: Cellulose + H2O →Glucose.
Fig. 2. Inhibition of cellulases: overview of the main kinetic pathways and interactions in
relation to the product inhibition: (a) main reactions (1, 2), (b) glucose inhibition (3, 4),
(c) cellobiose inhibition (5), (d) substrate inhibition (6, 7), (e) transglycosylation (8, 9).
amorphous and crystalline parts (Lynd et al., 2008; Mosier et al., 2005).
Nature has evolved certain cellulases to attack the crystalline cellulose, e.g.
the cellobiohydrolases, and others to attack the more amorphous regions,
notably the endoglucanases (Teeri et al., 1998). Nonetheless, this substrate
heterogeneity causes variations in the rates of the enzymatic hydrolysis of
the cellulose depending on the site of enzyme attack on the substrate. The
catalytic rates may also vary during the course of the cellulolytic
degradation since the cellulose which is hydrolysed at later stages of the
reaction is usually more crystalline than the (amorphous) cellulose
attacked during the initial stages of the reaction, and the degradation of
the crystalline cellulose is generally slower than that of the amorphous
parts (Lynd et al., 2002). As the reactions are heterogenous in nature, the
simple Michaelis–Menten equations do not suffice to describe these
reactions. The description of the kinetics of the actions of multiple
enzymes on insoluble, heterogenous lignocellulosic materials is in fact a
research subject in its own right and a number of kinetic models for
enzyme-catalyzed hydrolysis of cellulose have been proposed (e.g. Gan et
al., 2003; Kadam et al., 2004; Okazaki and Moo-Young, 1978; Wald et al.,
1984; Zhang and Lynd, 2004).
Despite the complexity of these dynamic models, the fundamental
base is of course a) that the enzyme must contact the substrate for a
reaction to occur and b) that an enzyme–substrate reaction
intermediate must form before the final product is released. Hence,
the model structures employed to describe the enzyme-catalyzed
hydrolysis of (ligno)cellulose are generally based on modifications of
Michaelis–Menten type kinetics. However, the models are often
combined with adsorption kinetics and may in some cases involve a
special “accessible substrate fraction” term (see Bansal et al., 2009;
Gan et al., 2003; Kadam et al., 2004; Zhang and Lynd, 2004). In
addition, inhibition terms are very frequently included to account for
the inhibitory actions caused by both cellobiose and glucose on
cellulolytic enzymes, and to a smaller extent the terms taking into
consideration the loss of enzyme activity for other reasons, e.g.
thermal inactivation and/or unproductive adsorption to lignin.
3.1. The Trichoderma reesei cellulase system
Most of the current knowledge about enzymatic cellulose hydrolysis stems from studies of the cellulolytic enzymes produced by
P. Andrić et al. / Biotechnology Advances 28 (2010) 308–324
T. reesei. The cellulolytic enzyme system of T. reesei has been
particularly extensively studied in relation to cellulose degradation
for bioethanol production as well as in relation to product inhibition
(as discussed later). The T. reesei cellulases have even been proclaimed
to be “the industry standard” for enzymatic lignocellulose hydrolysis
(Merino and Cherry, 2007). The enzymes secreted by T. reesei include
at least five different endoglucanases (EG I–V, or Cel7B, Cel5A, Cel12A,
Cel61A and Cel45A), two types of cellobiohydrolase activities (CBHI
(Cel7A) and CBHII (Cel6A)), as well as a number of xylanases and at
least one β-xylosidase enzyme (Reese et al., 1950; Rosgaard et al.,
2007b; Shoemaker et al., 1983). T. reesei also produces β-glucosidase
activity, but a large part of this activity is bound to the mycelium and is
therefore not efficiently recovered during industrial cellulase production from T. reesei (Rosgaard et al., 2007b). Although a β-glucosidase
from Aspergillus oryzae has been successfully expressed in a T. reesei
production strain (Merino and Cherry, 2007), a majority of application
studies employing T. reesei cellulases have up until now included
exogenous supplementation of a β-glucosidase preparation to boost
the β-glucosidase activity. This exogenous supplementation has often
been accomplished by addition of a β-glucosidase preparation from
Aspergillus niger (Andrić et al., 2010; García-Aparicio et al., 2006;
Jørgensen et al., 2007; Rosgaard et al., 2006, 2007a,b,c; Xiao et al.,
2004). The main purpose of this β-glucosidase addition has obviously
been to ensure the catalytic hydrolysis of cellobiose, and partly of the
cellulo-oligosaccharides, to glucose. However, as it will appear from
the discussion below, β-glucosidase addition may moreover alleviate
the product inhibition exerted by cellobiose on cellobiohydrolases and
endoglucanases.
3.2. Inhibition of cellulolytic enzymes by cellobiose and glucose
Cellobiose and glucose significantly decrease the cellulolytic
hydrolysis rate as well as the product yields in cellulose hydrolysis,
and both cellobiose and glucose are therefore to be considered as
inhibitors of cellulolytic enzymes (Gan et al., 2003; Gusakov et al.,
1987). Cellobiose, the product of cellobiohydrolase and partly of
endoglucananse action, directly inhibits both cellobiohydrolases
and endoglucanases (Gruno et al., 2004) (Fig. 2, path 5). Glucose
directly inhibits β-glucosidase (Dekker, 1986) (Fig. 2, path 3),
including the widely employed β-glucosidase from A. niger (Dekker,
1986; Xiao et al., 2004). Glucose also inhibits cellobiohydrolases
and endoglucanases directly (Holtzapple et al., 1990) (Fig. 2, path
4). In practice, however, the main inhibition by glucose on these
enzyme activities is an indirect inhibition because the glucose
inhibition of β-glucosidase may result in build-up of cellobiose. In
addition, cellulose and cellobiose, even being substrates, are also
known to exert some inhibition on cellobiohydrolases and
endoglucanases (Gong et al., 1977; Huang and Penner, 1991;
Väljamäe et al., 2001) as well as on β-glucosidase (Oh et al., 2000),
respectively (Fig. 2, paths 6 and 7). Several β-glucosidases,
including those from A. niger (Watanabe et al., 1992), T. reesei
(Schmid and Wandrey, 1989), and Thermonospora fusca (Ferchak
and Pye, 1983a) are furthermore able to catalyze a reverse reaction
in which glucose molecules via transglycosylation are transferred to
glucose or cellobiose to yield different di-, tri-, and oligosaccharides
(Fig. 2, paths 8 and 9).
4. Inhibition schemes
In classical Michaelis–Menten kinetics, encompassing simple
inhibition schemes, at least eight types of reversible inhibition have
been proposed: (i) competitive, (ii) uncompetitive, (iii) non-competitive, (iv) mixed, (v) partial, (vi) substrate, (vii) product, and (viii)
allosteric (Fullbrook, 1996). For cellulases, an inhibition type called
“hyperbolic” inhibition – a subtype of partial inhibition – has also been
suggested (Bezerra and Dias, 2004) (Table 1).
311
Table 1
Overview of common and special enzyme inhibition types. S (substrate), P (product),
I (inhibitor), E (enzyme). In relation to product inhibition of cellulases, all types from
the table are reported (I = P).
Common inhibition
Special inhibition
E + S ⇌ ES → E + P
Main reaction
Main reaction
E + S ⇌ ES → E + P
Type
Mechanism
Type
Mechanism
Competitivea
E + I ⇌ EI
Parabolicb
Uncompetitive
ES + I ⇌ ESI
Hyperbolicb
Mixedc/
Non-competitived
E + I ⇌ EI
ES + I ⇌ ESI
EI + S ⇌ ESI
Substratee
E + I ⇌ EI
EI + I ⇌ EII
E + I ⇌ EI
EI + S ⇌ ESI
EIS → EI + P
ES + I ⇌ EIS
ES + S ⇌ SES
a
Pseudo-competitive and partially competitive are also encountered within the
cellulase inhibition literature.
b
Rarely encountered.
c
Linear or complete inhibition is the most general case for common inhibition types.
d
Non-competitive inhibition is the special case of mixed inhibition in which the
inhibitor has the same affinity towards the free enzyme (E) and enzyme–substrate
complex (ES), and the substrate has the same affinity towards the free enzyme (E) and
enzyme–inhibitor complex (EI).
e
It is indirectly related to product inhibition of cellulases through e.g. cellulose or
cellobiose substrate inhibition.
4.1. Common inhibition kinetics
A competitive inhibitor lowers the substrate binding capacity and
increases the apparent KM because the enzyme–inhibitor complex
forms via direct interaction between the enzyme and the inhibitor
(Table 1).
In uncompetitive inhibition the inhibitor interacts with the
enzyme only when the enzyme is bound to its substrate (Table 1)
(Palmer, 1985). In mixed inhibition (Table 1), the inhibitor most often
binds to a remote site and induces a conformational change that
affects the active site (chemical properties or structure), reducing the
catalytic turnover and altering the substrate binding capacity (Pratt
and Cornely, 2004). In non-competitive inhibition the inhibitor is
visualized to form a non-productive complex with the enzyme irrespective of whether the enzyme is bound to the substrate or not
(Table 1) (Palmer, 1985; Pratt and Cornely, 2004).
For enzymatic cellulose hydrolysis a couple of special inhibition
schemes involving reactions of the enzyme:inhibitor complex (EI)
with either the inhibitor or the substrate have been reported (Bezerra
and Dias, 2004) — these rare schemes are usually designated as
parabolic and hyperbolic, respectively (Yoshino, 1987).
Substrate inhibition has been proposed to be a result of the inhibitory interaction between the enzyme–substrate complex (ES) and
the substrate (Fullbrook, 1996). Thus, in this case it appears that the
substrate at high concentration inhibits its own conversion to a
product (Table 1).
4.2. Product inhibition
Principally, the product of nearly every enzyme-catalyzed reaction
may behave as an inhibitor when present in high enough concentrations relative to the enzyme and substrate (Frieden and Walter, 1963).
In the simple Michaelis–Menten kinetic expression for the inhibited
reaction rate, v, (for the competitive type of inhibition), the inclusion
of the term I / KI, which is considered constant, accounts for this
inhibition effect:
v =
k E S
cat 0 KM 1 + KI + S
I
312
P. Andrić et al. / Biotechnology Advances 28 (2010) 308–324
(kcat is the turnover number, E0 is the initial enzyme concentration, S
is the substrate concentration, KM is the Michaelis–Menten constant, I
is the inhibitor concentration, and KI is the dissociation constant for
the enzyme:inhibitor complex).
In case of product inhibition however, the term I / KI in fact becomes
P / KI (P is product and KI is the dissociation constant for the inhibitory
enzyme:product complex). Since P increases with the progress of the
enzyme-catalyzed reaction, the term P / KI also increases and this
results in the catalyzed rate of reaction being reduced more and more
due to the product concentration increment. Since high product
concentrations are usually required in industrial processes (Fig. 1)
product inhibition may significantly retard reaction rates and hence
the efficiency of enzyme-catalyzed reactions at high extents of
substrate conversion in both batch and continuous processes.
Table 2
The special case of cellulose. Overview of enzyme adsorption and catalytic steps in
cellulase degradation of cellulose. E‐‐‐S enzyme adsorbed on cellulose, but not yet
reacted (non-complexed); ES′—non-productive enzyme–substrate complex; ES—
productive enzyme–substrate complex; ES*—enzyme–substrate complex activated
state; ESP*—enzyme–substrate–product complex activated state; E—free (soluble)
enzyme. Water is omitted as a reactant from the kinetic equations.
4.3. Diminishing the (product) inhibition effect
η=
Even though competitive inhibition of regular enzymatic reactions,
encompassing enzymatic attack on a soluble substrate, can be
outcompeted by substrate addition (Fig. 3), the case is different for
enzymatic reactions on insoluble substrates such as (ligno)cellulose.
With insoluble substrates, e.g. cellulose, the addition of more substrate
may not return the reaction rates to the uninhibited levels because
the slow-diffusing substrate entities are not able to compete for the
active site with fast-diffusing, soluble inhibitor molecules to form a
productive ES complex (Fig. 3). The inherently slower reaction rate of
the enzymatic action on the insoluble substrate (despite the presence
of a cellulose binding domain) may also be a factor (Holtzapple et al.,
1984a, 1990) (Fig. 3).
The principle of this problem is the same for a non-competitive
inhibitor, but for cellulases, the special case of reaction between the
inhibitor and the adsorbed, but not yet reacted enzyme (the latter
symbolized as [E‐‐‐S]), must be considered. Hence, the enzymatic
cellulose degradation includes a unique step where the enzyme is
adsorbed to the substrate, but where it has not (yet) reacted (Table 2).
As discussed in more depth later, the [E‐‐‐S] may progress to become a
productive ES complex, which progresses to the activated state, and
finally releases the product (Table 2). For processive enzymes such as
the cellobiohydrolases, the release of the product will not always result
in the liberation of the free enzyme, and the classical final step, written
as E + P, is rather [E‐‐‐S] + P. The [E‐‐‐S] then eventually progresses to
form ES and in turn yet another product molecule, still not necessarily
releasing the enzyme (Table 2). Alternatively, the [E‐‐‐S] may form a
non-productive complex ES′ (Table 2). In non-competitive inhibition
of cellulases, it may thus be envisaged that the inhibitor binds to the
free enzyme (E), the adsorbed, but not yet reacted enzyme ([E‐‐‐S]), or
the ES complex irrespective of the substrate presence (Table 1).
Holtzapple et al. (1984a) used a complexing (association)
constant, η, to describe the tendency of adsorbed, but not yet reacted
Main reaction: adsorption and catalysis
E + S ⇌ E‐‐‐S ⇌ ES( ⇌ ES * ⇌ ESP*) → E + P (productive)
E + S ⇌ E‐‐‐S ⇌ ES′ (non-productive)
ES → E‐‐‐S + P (for processive enzymes i.e. CBH)
cellulolytic enzyme ([E‐‐‐S]) to form the real enzyme substrate
complex (ES):
½E‐‐‐S
½ES
Two limiting cases of high and low complexing tendency (low and
high η, respectively), have been discussed elsewhere in relation to
diffusional limitations of the insoluble cellulose substrate, as the value
of η may determine the type of inhibition (Holtzapple et al., 1990).
Since the product inhibition challenges the efficiency of industrial
cellulolytic reactions it seems obvious and important to attempt to
alleviate it via reactor design that assures removal of the product(s) as
soon as they are formed.
5. Cellulolytic enzyme inhibition studies and results
Numerous attempts have been made to coin the experimentally
observed product inhibition mechanisms of cellobiose and glucose on
cellulases as competitive, non-competitive or mixed in accordance
with classical Michaelis–Menten schemes (Table 1). When examining
the available literature data on product inhibition of enzyme-catalyzed
degradation of cellulose by non-complexed, fungally derived enzyme
systems (presented in Tables 3 and 4), the competitive and noncompetitive types of inhibition have most commonly been proposed to
describe the inhibition exerted by glucose (Table 3) and cellobiose
(Table 4). Uncompetitive or even so-called pseudo- or partially
competitive inhibition mechanisms have also been suggested, however (Tables 3 and 4).
The investigations of cellulase inhibition by glucose and/or
cellobiose have mainly focused on: (1) Determination of the inhibition
mechanism and inhibition constant(s) on both initial rate measurements and extended reaction models and/or (2) Examination of the
inhibitory effect of product addition during extended reactions
(Tables 3–5). Unfortunately, the available experimental studies have
employed a variety of operation conditions of pH (4.5–6.5), temperature (38–60 °C), substrate dry matter content (1–10% (w/w)), initial
Fig. 3. Schematic representation of the enzyme action on soluble and insoluble substrates at a high level of externally added (soluble) inhibitor. The sketch is based on theory
presented by Holtzapple et al. (1984b, 1990). The second enzyme is cellulase with the characteristic binding domain, acting on insoluble cellulose, which is in turn composed of
glucose units (soluble inhibitor).
P. Andrić et al. / Biotechnology Advances 28 (2010) 308–324
313
Table 3
Overview of glucose inhibition constants and types for cellulase and β-glucosidase enzymes (the cellulose data are for cases with no β-glucosidase addition). Abbreviations: CMC:
carboxy methyl cellulose; PNPG: p-nitrophenyl-β-D-glucopyranoside; pNPβG: p-nitrophenyl-β-D-glucoside.
Enzyme source
KI [g/L]
Substrate
Inhibition type
Reference
Cellulase
T. reesei1
T. reesei2
T.reesei3, 4
T. reesei3, 4
T. reesei5
T. viride3, 6
T. viride7, 8
T. viride7, 9
T. longibrachiatum
Trichoderma3
A. niger10
Thermomonospora11
N/A9, 12
N/A8, 12
N/A13, 14
53
319
6
–
12
>9322
0.09–0.13
0.6
13
69
–
146
–
–
0.123
α-Cellulose
Dyed cellulose27
Solca Floc28
Solca Floc28
α-Cellulose
Cellulose29
Textile cotton/sulfate pulp
Textile cotton/sulfate pulp
Cellulose
–
CMC, wood shavings
Dyed cellulose27
Cellulose
Insoluble oligomers
Corn stover
Non-competitive
Non-competitive
Competitive
Uncompetitive30
Non-competitive
Non-competitive31
Uncompetitive
Competitive
Competitive
Non-competitive32
Non-competitive
Non-competitive
Competitive
Competitive
Competitive
Philippidis et al. (1993)
Holtzapple et al. (1990)
Asenjo (1983)33
Fan and Lee (1983)
Oh et al. (2000)
Gusakov and Sinitsyn (1992)34
Beltrame et al. (1984)
Beltrame et al. (1984)
Gusakov et al. (1985)
Tolan and Foody (1999)
Al-Zuhair et al. (2007)
Holtzapple et al. (1984b)
Mosier et al. (1999)
Mosier et al. (1999)
Kadam et al. (2004)34
0.6
0.1
0.0724
0.05–625
0.2–0.826
0.4
4–5
0.06
0.2–0.3
0.5
0.4
0.6
0.3
0.24
0.1–0.525
0.04–0.3
0.3
94
0.2–1.7
252
2.6–4
0.1–5
0.1–1.4
0.1–2.5
1.3
0.2
3.923
–
–
Cellobiose
Cellobiose
Cellobiose
Cellobiose
Cellobiose
Cellobiose
Cellobiose
–
PNPG
PNPG
PNPG
Cellobiose
Cellobiose
–
Cellobiose
Cellobiose
Cellobiose
PNPG
PNPG
pNPβG
Cellobiose
Cellobiose
–
–
Cellobiose
Cellobiose
Corn stover
Cellobiose
Cellobiose
Competitive
Competitive
Mixed
Competitive
Mixed30
Competitive
Non-competitive
Competitive
Competitive
Partially competitive
Competitive
Competitive
Non-competitive
Competitive
Competitve
Mixed
–
–
–
Competitive
Non-competitive
Competitive
Competitive
–
Competitive
Competitive
Competitive
Competitive
Non-competitive
Philippidis et al. (1993)
Asenjo (1983)
Wald et al. (1984)
Grous et al. (1985)
Gong et al. (1977)
Gusakov et al. (1992)
Hong et al. (1981)
Tolan and Foody (1999)
Yun et al. (2001)35
Dekker (1986)
Noble et al. (1990)
Beltrame et al. (1984)
Oh et al. (2000)
Tolan and Foody (1999)
Grous et al. (1985)
Alfani et al. (1990)
Gusakov et al. (1992)
Dekker et al. (2000)
Dekker et al. (2000)
Saha and Bothast (1996)
Noble et al. (1990)
Noble et al. (1990)
Saha and Bothast (1996)
Yun et al. (2001)
Lee and Fan (1983)
Fan and Lee (1983)
Kadam et al. (2004)34
Mosier et al. (1999)
Mosier et al. (1999)
β-Glucosidase
T. reesei1
T. reesei4, 7
T. reesei7, 15
T. reesei15
T. viride16
T. viride 6, 7
T. viride10
T. longibrachiatum
T. harzianum17, 18
A. niger19
A. niger19
A. niger3
A. niger19
A. niger
A. niger20
A. niger10
A. foetidus
A. foetidus17
5 Aspergillus strains17
Candida peltata17, 21
Various sources
Microbial enzymes17
Microbial enzymes17
N/A
N/A
N/A13
N/A12
N/A12
1
Laminex. 2Cellulase GC 123. 3Crude. 4QM9414. 5Celluclast 1.5 L. 6PBR (USSR). 7From cellulase. 8EG. 9CBH. 10Sigma-Aldrich. 11YX. 12Theoretical. 13From CPN. 14EG/CBH. 15Rutgers C30.16Isolated from comm. mixture. 17Purified. 18Type C-4. 19Novozym 188. 20Novo 250 L. 21NRRL (Y-6888). 22High BG activity. 23g/kg. 241.3 g/L for EG complex. 252 ranges of cellobiose
concentration. 262.6–2.9 g/L for EG complex. 27Azure dyed. 28SW 40. 29Modeling study. 30Called it non-competitive. 31Formally. 32Called it uncompetitive. 33From Lee YH, PhD thesis. 34From a
model. 35For a very tolerant KIG =54–252 g/L. 36QM9123 filtrate. 37BW200. 38Reactive orange dyed. 39Only due to analogy with glucose.
(added) glucose concentration (0.03–550 g/L), hydrolysis reaction time
(0.2–170 h); and have involved different substrate types and enzymes
from different sources — the latter moreover used in different
concentrations and reported in a range of units (Tables 3–5). Thus,
the comparison of the quantitative effects of inhibition by cellobiose
and glucose on cellulases is not straight forward.
With few exceptions (e.g. Andrić et al., 2010; Kadam et al., 2004)
only few quantitative data are available on inhibition of cellulases
acting on genuine, physico-chemically pre-treated, lignocellulosic
substrates — and even fewer have used high dry matter contents and
associated high glucose concentrations (Table 3). Rather, a large
number of cellulase inhibition studies have been accomplished using
different “pure” cellulose substrates such as Solca Floc, Avicel, CMC,
and α-cellulose (Table 3).
Despite the significant variations in the experimental conditions
employed, and the ambiguities and shortcomings in the mathematical
descriptions of the enzyme-catalyzed events and the inhibition
mechanisms of cellulases (discussed later), the data nevertheless
signify that product inhibition profoundly decreases the rates of
enzyme-catalyzed cellulose hydrolysis (Table 5). Cellobiose exerts the
strongest inhibition effect on cellulase activity with typical KI ranges
between 0.01 and 6 g/L (Table 4). Glucose exerts a similar, strong
inhibition on the activity of β-glucosidases with KI typically ranging
from 0.04–5 g/L (Table 3). Glucose also inhibits the cellulase activity
(other than the β-glucosidase activity), but to a lower extent, with KI
ranges typically varying from 0.1–70 g/L (Table 3).
5.1. Glucose inhibition of cellulolytic enzymes
The reported extents of the decrease in reaction rate induced by
glucose inhibition vary from ∼10–100% depending on the system set-up
with respect to the substrate, the enzyme:inhibitor levels, the extent of
reaction at which the inhibition was assessed, and not least the enzyme
system employed (Table 5). The available data confirm that the
inhibition exerted by glucose varies significantly in different systems
(different substrates, enzyme:substrate dosage levels, glucose levels,
314
P. Andrić et al. / Biotechnology Advances 28 (2010) 308–324
Table 4
Overview of cellobiose inhibition constants and types for cellulases enzymes (legends explanation is given in Table 3).
Enzyme source
KI [g/L]
Substrate
Inhibition type
Reference
T. reesei1
T. reesei2
T. reesei3
T. reesei5
T. reesei15
T. reesei4, 8, 17
T. reesei4, 9, 17
T. reesei9
T. reesei5
T. reesei9, 17
T. reesei.3, 4
T. viride3, 8
T. viride3, 9
T. viride3, 6
T. viride36
Trichoderma3
T. longibrachiatum
A. niger10
Thermomonospora11
N/A13, 14
N/A9, 12
5.9
54
–
6.0
0.05
4–12
0.5
0.01
0.02
0.01
2.3
0.4
1.8
3.6
1.6
3.3
4.2
–
11
0.02
–
α-Cellulose
Dyed cellulose27
Solca Floc28
α-Cellulose
Rice straw
Amorphous cellulose
Bacterial cellulose
Avicel
Avicel
Avicel
Solca Floc28
Sulfate pulp
Sulfate pulp
Cellolignin
Solca Floc37
–
Cellulose38
CMC
Dyed cellulose27
Corn stover
Cellulose
Non-competitive
Non-competitive
Uncompetitive30
Non-competitive
Uncompetitive30
–
General mixed
Competitive
Non-competitive
Competitive
Competitive
Uncompetitive
Competitive39
Uncompetitive
Non-competitive
Non-competitive32
Competitive
Non-competitive
Non-competitive
Competitive
Competitive
Philippidis et al. (1993)
Holtzapple et al. (1990)
Fan and Lee (1983)
Oh et al. (2000)
Wald et al. (1984)
Gruno et al. (2004)
Gruno et al. (2004)
Bezerra and Dias (2004)
Bezerra and Dias (2005)
Bezerra and Dias (2005)
Asenjo (1983)33
Beltrame et al. (1984)
Beltrame et al. (1984)
Gusakov et al. (1992)
Howell and Stuck (1975)
Tolan and Foody (1999)
Gusakov et al. (1985)
Al-Zuhair et al. (2007)
Holtzapple et al. (1984b)
Kadam et al. (2004)34
Mosier et al. (1999)
1
Laminex. 2Cellulase GC 123. 3Crude. 4QM9414. 5Celluclast 1.5 L. 6PBR (USSR). 7From cellulase. 8EG. 9CBH. 10Sigma-Aldrich. 11YX. 12Theoretical. 13From CPN. 14EG/CBH. 15Rutgers C30.16Isolated from comm. mixture. 17Purified. 18Type C-4. 19Novozym 188. 20Novo 250 L. 21NRRL (Y-6888). 22High BG activity. 23g/kg. 241.3 g/L for EG complex. 252 ranges of cellobiose
concentration. 262.6–2.9 g/L for EG complex. 27Azure dyed. 28SW 40. 29Modeling study. 30Called it non-competitive. 31Formally. 32Called it uncompetitive. 33From Lee YH, PhD thesis. 34From a
model. 35For a very tolerant KIG =54–252 g/L. 36QM9123 filtrate. 37BW200. 38Reactive orange dyed. 39Only due to analogy with glucose.
etc.). Although definite conclusions cannot be made because of the large
variation in the data material the T. viride cellulases seem to be more
sensitive to glucose inhibition than the T. reesei cellulases (Table 3).
The inhibition of cellulases is complicated to assess because, as
shown by Gusakov and Sinitsyn (1992), the efficiency of a cellulolytic
enzyme mixture in converting cellulose to glucose, and consequently
Table 5
Overview of the studies investigating the effect of glucose addition on enzymatic hydrolysis of (ligno)celullose.
Enzyme source
Substrate
S0 [% w/v]
Added glucose [g/L]
Time [h]
Cellulase
T. reeseia
T. reeseie
T. reeseih
T. reeseim, n
T. reeseim, n
T. reeseit
T. reeseiv
T. longibrachiatumw
Thermomonosporay
Thermomonospora fuscay, aa
Clostridium thermocellumad, ae
Penicillium funiculosumai
N/Aak
Clostridium thermocellumaa
Clostridium thermocellumaa
Dyed cellulose
α-Cellulose
Rice strawi
Solca Floco
Solca Floco
Avicel cellulose
α-Cellulose
Dyed cellulose
Dyed cellulose
Swollen celluloseab
CMC
Solca Floco
Corn stoveral
Swollen Avicel
Microcrystalline Avicel
0.5
6
10j
–
5j
2/10
9
4
0.1–1
2–4
0.5
5
10j
0.06
0.06
546
20/80
20
30
10/30
20
10/80
20
100/200
200
0.03
30
30/50am
50
60
0.25–1b, c,
0.02f, c, g
47k, l
8k, p
8q, r, s
0.5k, c, r, u
−f, c
1xk, b, f, x
1b, c, zx
4f, ac
20ag, ah
8q, aj
168k, an
3ah
18ah
Cellulase and β-glucosidase
T. reeseiao/A. nigerap
T. reeseiv/A. nigerap
T. reeseit/A. nigerap
Solca Flocaq
Wheat strawat
Sallowaw, ax
10j
2
10
50/100
20/40
25/50
β-Glucosidase
T. reeseie
A. nigerap
A. nigerap
A. nigerap
A. nigerap
A. nigerap
Thermomonospora fuscay, ba
Thermomonospora fuscay, ba
Candida peltatabc
15 yeast strains
Cellobiose
Cellobiose
Cellobiose
Cellobiose
PNPG
Cellobiose
PNPG
Cellobiose
Cellobiose
–
1
1
1
2/10
0.15
1
–
1
10
–
3/20
120
50/100
20/100
20
10/20–45
200
150
60
300
, af
Inhibition effect [%]
Reference
63–66
20/60
<5–7
46
10/44
55/35
55/78
48/15/0
29/58
60
50
67
15/20
10
35
Holtzapple et al. (1990)
Philippidis et al. (1993)
Wald et al. (1984)
Asenjo (1983)
Lee and Fan (1983)
Xiao et al. (2004)
Oh et al. (2000)
Gusakov et al. (1985)
Holtzapple et al. (1984b)
Ferchak and Pye (1983b)
Mosolova et al. (1993)
Rao et al. (1989)
Kadam et al. (2004)
Johnson et al. (1982)
Johnson et al. (1982)
4–24q, ar, as
96k, au, av
72as, ay
23–18/60–37
46/62
16/38
Tjerneld et al. (1985)
Andrić et al. (2010)
Frenneson et al. (1985)
0.02k, c, g
0.5k, au
6k, au
0.5k, c, r, az
0.5k, c, r, az
−k, c
−ac, bb
4k, ac
144k, bd
–
75/95
63
10/25
47/52
90
75/100
65
30
≈0
<50
Philippidis et al. (1993)
Dekker (1986)
Dekker (1986)
Xiao et al. (2004)
Xiao et al. (2004)
Oh et al. (2000)
Ferchak and Pye (1983a)
Ferchak and Pye (1983a)
Saha and Bothast (1996)
Saha and Bothast (1996)
d
a
Cellulase GC 123. bDye release. cInitial rate. dpH 4.5, 49 °C. eLaminex. fgpH 5, 38 °C, 25 IU/gcellulose. hRutgers C30. iHammer-milled, acid treated. jkGlucose release. lpH 5, 45 °C, 2 FPU/
mL. mnQM9414.oSW 40. pLee YH, PhD thesis. qReducing sugar release. rpH 4.8, 50 °C.s67% based on initial rate. tu40 FPU and 80 CBU/gcellulose. vCelluclast 1.5 L. wEthanol precipitated
preparation. xpH 4.5, 50 °C, 0.28 FPU/mL. yYX.z55 °C. aaCulture filtrate. abThe same on 5% Solca F. acpH 6.5, 55 °C. adIsolated. aeExpressed in Escherichia coli. afEG. agViscosity reduction.
ah
pH 6, 60 °C. aiNational Chemical Laboratory India. ajpH 4.8, 50 °C, 6.5 FPU/gsubstrate. akCPN.alDilute-acid pre-treated. amg/kg. anpH 4.8, 45 °C, 15 FPU/gcellulose. aoCellclast 2.0 L Type X.
ap
Novozym 188. aqBW 200. arpH 4.8, 50 °C, 1.4 FPU/mL. asCellulose conversion. atHydrothermally pre-treated. aupH 5, 50 °C. av8 FPU/gDM, 13 CBU/gDM. awHammer milled; NaOH
bc
thestandard
inhibition
depend
heavily
on the concentration and
treated. axQ082. aypH 4.8, 40 °C, 1.4 FPU/mL. az80 CBU/gcellobiose/PNPG. baGeneral Electric. bbPNPG
assay. pattern,
Purified. bd
pH 5, 50 °C,
1.5 U/mL.
P. Andrić et al. / Biotechnology Advances 28 (2010) 308–324
activity of the present β-glucosidase enzymes — in addition to the
other factors (Fig. 2). The main problem is, however, that the
“fundamental” interpretation of multi-enzyme-catalyzed cellulose
hydrolysis reactions, and hence their inhibition, rests on solving an
equation with at least two unknown quantities: the heterogenous
substrate and the enzyme mixture. In addition, as for other enzymes,
the extent of inhibition will first and foremost depend on the relative
levels of the inhibitor(s), in this case both glucose and cellobiose, in
relation to the concentrations of the enzymes. Although the value of
the KI may vary for different cellulases and cellulase systems, the KI
for a competitive inhibition reaction should in theory not depend on
substrate properties, and the KI for mixed inhibition may only
indirectly depend on substrate properties (Table 1). In any case,
however, KI should theoretically resemble the enzyme–product
interaction, also within the ESP complex, which can be formed only
if the: (a) enzyme has another site for product attachment, or (b)
catalytic site has multiple binding subsites. The apparent overall KM
(Michaelis–Menten constant) value for cellulolytic reactions presumably changes with the reaction progress and this change in KM
indirectly contributes to the variation in the reported KI values. Hence,
it is clear that the different model structures and hydrolysis schemes
must result in different parameter values and make direct comparisons of different models difficult (Bansal et al., 2009; Kadam et al.,
2004). The fact that the reported values of the KI for glucose inhibition
of cellulases range from 0.1 to 319 g/L (Table 3) indicates that the
currently employed inhibition models may in fact be quite empirical.
5.2. Glucose inhibition of β-glucosidases
The product inhibition exerted by glucose on various fungal βglucosidases has mainly been reported to take place via a competitive
inhibition mechanism — although other types of inhibition, notably
non-competitive and mixed inhibition have also been proposed
(Table 3). In contrast to the complexity of the coordinated action of
endoglucanases and cellobiohydrolases on insoluble, heterogenous
cellulose, which complicates the modeling and the understanding of
the entire enzymatic hydrolysis reaction, the β-glucosidase catalyzed
cellobiose hydrolysis and the product inhibition of β-glucosidase
exerted by glucose may indeed follow Michaelis–Menten kinetics.
This is because the cellobiose is soluble and because the hydrolysis of
this substrate takes place via a simple “one-attack” hydrolysis reaction
(Andrić et al., 2010; Grous et al., 1985). In this light, the proposed
competitive product inhibition exerted by glucose on β-glucosidase
may indeed be the most plausible inhibition mechanism. Except in a
few cases, e.g. notably the data obtained using p-nitrophenolsubstrates (Table 3), the reported KI values for glucose inhibition of
β-glucosidase activity are quite consistent and range from ∼ 0.1 to
0.8 g/L with no particular differences in the KI values between the most
studied T. reesei and A. niger β-glucosidase activities. The trend in the
data is that the T. reesei β-glucosidase activity is more prone to
inhibition than the A. niger and the T. viride β-glucosidases, (Table 3);
however, as the β-glucosidase activity present in the cellulase
preparation or externally supplemented may vary widely (see
below), this trend may not necessarily constitute a definite predisposition. A few significantly different KI values for glucose have been
reported for other β-glucosidase enzymes. The reported KI value for a
β-glucosidase from Candida peltata, as assessed on p-nitrophenyl-β-Dglucoside, was for example 252 g/L, while that of Aspergillus foetidus βglucosidase in one case was reported to be 94 g/L — again on pnitrophenyl-β-D-glucoside as substrate (both highly glucose tolerant,
Table 3). In another study the KI value for an A. foetidus β-glucosidase
was found to be 0.3 g/L assessed on cellobiose (Gusakov et al., 1992)
(Table 3). Although the highest and seemingly most extreme KI
data have been obtained on artificial p-nitrophenyl-β-D-glucoside
substrates, there are no obvious kinetic or mechanistic artifacts that
should result from the use of these substrates and which then would
315
offer an explanation to the high KI values reported. The p-nitrophenylβ-D-glucoside substrates have also been employed in studies where the
KI values obtained were in the order of 0.2–1.7 g/L (Table 3). In applied
cellulose hydrolysis, including glucose inhibition studies, β-glucosidase is often added in surplus to boost the glucose production e.g. with
T. reesei cellulases (see e.g. Andrić et al., 2010; Kadam et al., 2004).
In these reactions the overall reaction kinetics may be highly affected
by the β-glucosidase kinetics, and the modeling of the inhibition
kinetics may in turn fit classical Michaelis–Menten models surprisingly well (Andrić et al., 2010).
5.3. Inhibition by cellobiose
Several of the published studies on glucose inhibition of cellulases
have also evaluated the inhibition exerted by cellobiose on different
(non-complexed) cellulolytic enzyme systems. The mechanism of
cellobiose product inhibition has been described as non-competitive,
uncompetitive, competitive, and even as “mixed” inhibition (Table 4),
and the reported values for the inhibition constant, KI (g/L), vary
widely: from 0.01 to 54 g/L. However, the right value, under typical
reaction conditions, may rather be in the order of 3–6 g/L (Table 4). In
a modeling study examining the actions of T. viride cellulases on
“pure” cellulose, both non-competitive, pseudo-competitive, uncompetitive and mixed inhibition models were shown to describe the
cellobiose product inhibition (Gusakov and Sinitsyn, 1992) (Table 4).
In a more recent modeling study from The US National Renewable
Energy Laboratory (NREL), with corn stover as the lignocellulosic
substrate, the cellobiose inhibition was proposed to be competitive
(Kadam et al., 2004).
There is no trend in the available data that indicate whether the
effect of cellobiose inhibition on cellulases differs for particular types
of substrates in response to e.g. crystallinity of the substrate (Table 4).
The variation in the data obtained on different substrates could also be
related to the differences in the estimation of model parameters used
in different reports.
As discussed later, the inconsistencies in the reported types of
inhibition and in the reported KI values may – at least partly – be a
result of the complex kinetics of enzymatic cellulose degradation and
a consequence of significant differences in experimental conditions
employed in different studies. However, even for the very many
studies assessing the cellobiose inhibition kinetics on the T. reesei
cellulases – on various more or less “pure” cellulose substrates – there
are significant inconsistencies in the types of inhibition reported for
the inhibitory action of cellobiose, and in the values determined for
KI (Table 4). Cellobiose is the direct product of the actions of
cellobiohydrolases, but may also “accidentally” result from endoglucanase activity (Fig. 2). T. reesei produces two distinct cellobiohydrolases, CBHI and CBHII, respectively, that attack at the reducing and
non-reducing ends of cellulose chains, respectively (Teeri et al., 1998).
The significance of the cellobiohydrolase action for the cellulose
degradation depends on the crystallinity of the substrate. Among
the different cellulase activities, cellobiohydrolases have the
highest apparent activities on crystalline cellulose. The substrate
crystallinity changes during the enzymatic hydrolysis with mixed
cellulases with the amorphous parts usually being hydrolysed first.
The impact of cellobiose inhibition on cellobiohydrolases will
therefore change depending on the substrate crystallinity and may
in turn vary with degree of hydrolysis. In conclusion, the inhibition
data and KI obtained in different studies will vary unless the degree of
substrate crystallinity and the composition of the cellulase mixture
have been the same. The inconsistencies in the reported KI values
simply reflect that this may not have been the case. The inconsistencies
in the reported KI values clearly indicate the challenges of conducting
and interpreting kinetic studies, including inhibition studies, on
enzyme-catalyzed cellulose hydrolysis (discussed further in Section 7,
below).
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5.4. Cellulase inhibition terminology controversies
Even though the kinetics and the type(s) of inhibition mechanisms
exerted by cellobiose and notably by glucose on different cellulolytic
enzymes have been extensively studied (Tables 3 and 4) there is a
surprising lack of a definite agreement on exactly how the overall
reaction and how the action of the different cellulolytic enzymes are
retarded by cellobiose and glucose. One of the primary inconsistencies
in the reported mechanisms for cellulase product inhibition may likely
be due to the prerequisite for the cellulolytic hydrolysis reaction,
namely the adsorption of cellulases (cellobiohydrolases or endoglucanases) on the available or rather the accessible sites on the insoluble
cellulosic substrate. This is opposed to the catalytic rate being a direct
function of the gross cellulose substrate concentration. Although the
available substrate is related to the total substrate level, the accessible
substrate concentration is in most cases unknown. In addition, the
further challenge in understanding the exact inhibition mechanism is
that the enzymatic degradation of cellulose indeed requires the
concerted action of several enzymes, which may be subject to
differentiated inhibitor binding and inhibition mechanisms.
Finally, some confusion has also arisen due to the inconsistent use of
inhibition kinetics terminology. Lee and Fan (1983) originally proposed
an “uncompetitive” type of inhibition mechanism for the inhibition
exerted by the products, i.e. cellobiose and glucose, on cellulases
(Table 1). However, in their own subsequent publication they termed
this inhibition as “non-competitive” (Fan and Lee, 1983). Wald et al.
(1984) described the mechanism of inhibition of cellulases to be
“uncompetitive” (according to Table 1), but termed it “non-competitive”.
Later, Gusakov and Sinitsyn (1992) claimed that the model of inhibition
originally proposed by Lee and Fan (1983) should be classified as “noncompetitive”. In their model Gusakov and Sinitsyn (1992) proposed that
the cellobiose inhibits the cellulases via interactions taking place between
cellobiose and cellulases adsorbed to the substrate — a model which
kinetically resembles an “uncompetitive” inhibition mechanism according to classical terms (Table 1). Zhang and Lynd (2004) classified the
mechanism originally proposed by Lee and Fan (1983) as “uncompetitive” and the one from Gusakov and Sinitsyn (1992) as “noncompetitive” even though the two mechanisms appear to be identical.
In complete contrast to this, Tolan and Foody (1999) used the term
“uncompetitive” to describe what is essentially a rate expression for a
“non-competitive” inhibition. Other inhibition type terms have been
ambiguously used as well. Gong et al. (1977) for example used the
term “simple non-competitive” to designate the non-competitive
inhibition, while the mixed inhibition was termed as “non-competitive”; Oh et al. (2000), on the other hand, described non-competitive
inhibition of β-glucosidases but presented a rate equation for the
competitive type of inhibition.
Whether it is necessary to strictly distinguish between these
inhibitions mechanisms in genuine processing of lignocellulose is
actually not certain. Nonetheless it seems clear that significant
confusion has arisen from the ambiguous use of terminology, especially
the varied use of “non-competitive” and “uncompetitive” inhibition.
5.5. Mathematical curiosity of non-competitive modeling of cellulolytic
inhibition
For the enzyme-catalyzed degradation of straw lignocellulose with
the T. reesei cellulase and the A. niger β-glucosidase system (i.e. the
widely used Celluclast 1.5 L + Novozym 188 system, Novozymes A/S
Bagsværd, Denmark) we recently observed that the product inhibition
exerted by glucose on this multi-enzymatic reaction was surprisingly
well described by simple Michaelis–Menten inhibition models
(Andrić et al., 2010). Although the significance of the inhibition
mechanism on the quality of the fit in extended reactions was in fact
small, we observed that the non-competitive inhibition mechanism
fitted the glucose inhibition data the best (Andrić et al., 2010). The
question is then, also in relation to the results reported in Table 3, why
this product inhibition, which in reality would most likely be a
competitive product inhibition, may be best described as a noncompetitive type of inhibition?
The non-competitive inhibition mechanism assumes that the
inhibitor, in this case glucose, interacts with both the enzyme and
the enzyme substrate complex (Table 1). In the simplified rate
expression describing this non-competitive inhibition, the dissociation constant for the inhibitor, i.e. glucose, and the enzyme (KI) and
that of the inhibitor and the enzyme–substrate complex (KI’) are
assumed to be equal, and described as KI. In effect, this expression then
results in a multiplication factor influence of the P / KI “inhibition
factor” (P is glucose) on both the substrate concentration (S) and the
enzyme–cellulose complex dissociation constant, KM , which affects
the rate, v, negatively (Andrić et al., 2010):
v =
kcat E0 S
ðKM + SÞ 1 +
P
KI
(kcat is the turnover number, E0 is the initial enzyme concentration, S
is the substrate concentration, KM is the Michaelis–Menten constant,
P is the product inhibitor concentration, and KI is the dissociation
constant for the enzyme:inhibitor complex).
Hence, as the only one amongst the Michaelis–Menten inhibition
models, this non-competitive model incorporates an additional
inhibitory effect to be multiplied with the substrate concentration
“on top of” the direct competitive product inhibition by glucose,
which only affects the KM because the denominator in the Michaelis–
Menten model for competitive inhibition is S + KM (1 + I / KI).
It is likely that the inclusion of such an extra inhibitory effect in the
Michaelis–Menten reaction rate expression may partly adjust for
some of the complex influence from the substrate or perhaps from the
multi-enzymatic cellulose degradation rather than defining a true
non-competitive inhibition mechanism.
6. Experimental problems of enzymatic cellulose hydrolysis in
relation to understanding product inhibition
Although it immediately appears a relatively practicable task to
accomplish fundamental studies on industrially relevant cellulose
conversion reactions, a number of conceptual and experimental
challenges exist. These challenges have likely affected the results
obtained up until now (Tables 3–5). As outlined in the following, the
main challenges are related to: a) the measurement of the product
inhibition rates, b) the complexity of the enzymatic accessibility on the
cellulosic substrate, and c) the kinetics of the multiple enzymes attack.
Recognition of the experimental challenges of enzymatic cellulose
hydrolysis in relation to assessing and understanding product inhibition
will hopefully provide for improved experimental protocols in the future.
6.1. Product inhibition study strategy
A general problem in the examination of product inhibited
enzymatic reactions is that the rate measurements and data interpretation are complicated by the fact that the inhibitor concentration
changes as the reaction progresses. During extended cellulolytic
hydrolysis reactions an additional issue is the time factor because
the cellulolytic hydrolysis is in fact slow. Hence, the cellulolytic
enzyme protein molecules may be subject to inactivation e.g. because
of shear or thermal inactivation during the time of reaction (Gunjikar
et al., 2001; Rosgaard et al., 2007c). This inactivation in turn influences
the cellulose hydrolysis rates and it is difficult to separate the
assessment of this effect from the effects from inhibitors.
In order to assess the inhibition exerted by the reaction products
cellobiose and glucose, these products may be deliberately added to the
reaction mixture to measure their direct effect on the reaction rate — as
P. Andrić et al. / Biotechnology Advances 28 (2010) 308–324
discussed in more depth later, most of the available product inhibition
studies on cellulases have in fact employed this strategy: in typical
batch-reactor cellulase inhibition studies, the inhibitor, typically
glucose, has been added to result in a relatively high concentration,
e.g. ≥20 g/L, as compared to the substrate level (Andrić et al., 2010).
These high (inhibitory) product levels result in low substrate-toproduct (S/P) and low enzyme-to-product (E/P) ratios, if assuming that
E is constant. Under genuine conditions of the enzymatic hydrolysis, i.e.
without glucose added, the S/P and E/P ratios will however be relatively
high during the major part of the reaction, as the present glucose will
originate only from the reaction and then build up to a significant
concentration only during the reaction.
Another complication is that the reaction rate of cellulose hydrolysis
is usually measured by assessing the rate of product formation — as
opposed to directly measuring the rate of substrate consumption.
Hence, when glucose is added, the measurement of the products formed
over time, which defines the rate, must be done on a background of the
added levels of that product. Even with modern analytical methods the
measurement of small changes on a high background inherently
decreases the sensitivity of the measurements. This problem has been
attempted avoided by use of model substrates, i.e. cellulosic materials
onto which a dye or a radioactive group has been attached, e.g. azur
cellulose or reactive orange cellulose or radioactive reducing-end
labeled cellulose (Gusakov et al., 1985; Gruno et al., 2004; Holtzapple
et al., 1984b, 1990). Although the hydrolysis of these substrates allows
the discrimination between added glucose and the product(s) released
as a result of the substrate hydrolysis, their drawback is that the
enzymatic attack and hydrolysis rates may not be similar to those on
unmodified substrates. Secondly, if the initial rates or yields of
hydrolysis (conversion degree) are measured as the release of dye or
radioactively labeled reducing sugars the practical conduction of
experiments may become very complicated.
With this in mind, an optimal solution would be to assess the rate
in the presence of glucose, e.g. in a batch reactor, and then compare it
to the rate recorded in a reactor in which the glucose was instantly
and/or completely removed from the reactor (Alfani et al., 1990).
6.2. The cellulosic substrate
Cellulose encompasses regions of relatively highly ordered
polymeric, crystalline molecules protected inside micro-fibrils. In
lignocellulose-to-ethanol processing, the pre-treatment step of
lignocellulose is assumed to produce an increased level of amorphous
regions that are interwoven with the crystalline cellulose (Mosier
et al., 2005), but some cellulose micro-fibrils stay intact, i.e. apparently
retaining a relatively high degree of crystallinity even after extensive
pre-treatment (Kristensen et al., 2008b; Pedersen and Meyer, 2009).
Although fungal cellobiohydrolases actually exhibit activity on crystalline cellulose (Teeri et al., 1998), the crystalline regions are inherently
more resistant to the cellulolytic enzymes attack than the amorphous
regions (Teeri et al., 1998; Zhang and Lynd, 2004). On top of this, the
lignin presence can both act as a sterical hindrance and as a reversible
(in practice: irreversible) enzyme inhibitor, reducing the efficiency
of the cellulases (Chang and Holtzapple, 2000).
As already mentioned above, the majority of the classical cellulase
inhibition studies have employed model cellulosic substrates like
Solca Floc, α-cellulose, Avicel, bacterial and amorphous cellulose,
cotton, and more rarely soluble cellulose derivates like CMC (Tables 3
and 4). Product inhibition including determination of the kinetic
mechanism and the KI has been quantified from both initial rate and
extended reaction measurements. Obviously, because of the physical
and chemical differences of the substrates with respect to crystallinity, degree of cellulose polymerization etc., the data obtained have
varied widely (Table 3).
In relation to industrial processing, it might seem more relevant to
employ genuine pre-treated lignocellulosic substrates, e.g. corn stover
317
or wheat straw (Kadam et al., 2004). However, with genuine substrates,
the composition and the levels of accessible substrate vary significantly,
and it is actually not possible to obtain universally valid data. In order to
at least obtain comparable data for inhibition of cellulolytic enzymes
among different laboratories, e.g. when assessing novel enzymes, it
would be optimal if an agreement was made in the global scientific
community, that a universal cellulosic model substrate, with standardized physical properties was used. The substrate should be standardized
with respect to chemical composition, degree of crystallinity, degree
of polymerization, solubility, and included in cellulase inhibition studies
at least as a “control”. If such an agreement was made, e.g. as a
requirement for publication of data, it might also be relevant to develop
a particular protocol for the enzymatic reaction with respect to
employing a certain level of dry matter, a certain temperature, a certain
reaction time, a certain enzyme protein dosage etc., as similar to the
current filter paper unit (FPU) assay — in fact, it may be a part of a new
‘extended’ FPU assay!
6.3. The cellulolytic enzymes
Although several studies have employed purified or cloned monocomponent cellulases, cellobiohydrolase, endoglucanase or β-glucosidases, different multicomponent cellulase preparations from Trichoderma spp. (notably T. reesei, T. viride, T. longibrachiatum and
T. harzianum) have most commonly been employed in cellulase inhibition studies (Tables 3 and 5). The enzyme activities and employed
dosage levels have varied significantly: enzyme addition levels have
ranged from 0.28 to 2 Filter Paper Units (FPU)/mL equivalent to roughly
12–40 FPU/gcellulose (Table 5). Due to the complexity of the cellulolytic
enzyme mixture having individual components that can behave rather
differently (depending on origin, nature, composition and multiplicity),
the current data do not permit finite conclusions to be drawn with
respect to the glucose and cellobiose inhibition effects towards different
enzymes from different sources (Tables 3–5). As already mentioned,
the cellulase mixtures being deficient in cellobiase activity, i.e. notably
the T. reesei cellulases, have been supplemented with the extra βglucosidases from A. niger or A. foetidus — typical addition levels range
from 25 to 80 cellobiase units/gcellulose. The β-glucosidase activities from
certain yeast strains, including those from Candida sp. (Saha and
Bothast, 1996 and Table 5) have been reported to be relatively robust
to glucose inhibition during cellobiose hydrolysis. In general, however,
the available data do not provide any clear conclusions as to whether
certain cellulolytic enzyme systems are more resistant than others to
glucose inhibition (Table 5). The current data clearly signify, however,
that the widely used T. reesei enzymes are significantly inhibited by
glucose (Tables 3 and 5).
The difficulty in the comparison of the effect of inhibiting products on
cellulase enzymes, in a multi-enzyme and thus multi-step reaction with
intermediate products, is found primarily in a number of combinations
of enzymes, enzymes sources/forms, activities that are desired to
evaluate, methods for inhibition evaluation and inhibitors (cellobiose,
and glucose) on which the inhibition is tested (Table 6). The analytical
tools for product determination may also be important as in very many
cases reducing sugars have been reported, making it tough to
understand the effect of a product inhibitor on a specific enzyme activity.
Cellulases are naturally a battery composed of endoglucanase,
cellobiohydrolase and β-glucosidase enzymes, but depending on
the form of enzyme production, they may have quite a different level
of β-glucosidase activity. The cellulase degrading effect on cellulose
may be tested by observing the reaction to cellobiose as a product (in
an enzyme mixture of low β-glucosidase activity) or to glucose (higher
β-glucosidase activity); in both cases either cellobiose or glucose
may be added as inhibitors, depending on the research goal (Table 6).
Cellulases may further be investigated for product inhibition on its
β-glucosidase activity in which case the inhibiting glucose is added to
a mixture containing substrate cellobiose and a whole battery of
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P. Andrić et al. / Biotechnology Advances 28 (2010) 308–324
Table 6
Summary of research strategies for performing cellulase inhibition study. RS — reducing sugars.
Enzyme
Enzyme form
Enzyme activity
to evaluate
Inhibition evaluation methoda
Inhibitor addition
Substrate
Cellulaseb
Crude /purified/commercial
Cellulase + β-glucosidasec
Crude /purified/commercial
β-glucosidase
Crude /purified/commercial
Cellulase
β-Glucosidaseb
Cellulase
β-Glucosidased
β-Glucosidase
RS/cellobiose/glucose release
Glucose release
RS/cellobiose/glucose release
Glucose release
Glucose release
Cellobiose/glucose
Glucose
Cellobiose/glucose
Glucose
Glucose
Cellulose
Cellobiose
Cellulose
Cellobiose
Cellobiose
a
b
c
d
Other methods are described in the text.
β-Glucosidases are natural part of the cellulase mixture.
Typically commercial enzymes.
Not seen in practice.
cellulases (Table 6). Theoretically, this may be done with cellulases that
are supplemented with β-glucosidase activity. This is not seen in
practice, however, rather, in this case, the glucose inhibition is studied as
on a pure β-glucosidase enzyme (Table 6). In any case, the overall
cellulase performance will be heavily influenced by the presence of
β-glucosidase activity that additionally indirectly affects the performance of other enzymes in the mixture.
In order to compare the inhibition data of cellulolytic enzymes
from different preparations it would in theory be necessary to
introduce designed mixtures of purified or cloned mono-component
cellulase enzymes and test the product inhibitory effect for each
enzyme separately. However, the main problem is of course that the
outcome of the action of the individual activities may neither be easy
nor relevant to measure, because the degradation of cellulose to
glucose obviously requires cooperative action of several enzymes.
If the purpose of the study is to quantify the inhibition involving
the determination of the degree of inhibition exerted by the products
in order to predict the hydrolysis extent at longer reaction times — the
strategy would be to use a defined cellulase mixture. If the process
requirement is the maximization of the glucose production the right
strategy might be addition of sufficient β-glucosidase activity to
catalyze the complete conversion of cellobiose to glucose and to
minimize the negative effect of cellobiose.
6.4. Enzyme-to-substrate ratio (E/S)
As shown by Gusakov and Sinitsyn (1992), the E/S ratio is also a
crucial parameter to consider when investigating cellulase product
inhibition and kinetics. If E/S is relatively high, the cellulose surface – i.e.
the reactive sites on the substrate – may be “saturated” with adsorbed
enzymes and a fraction may even remain in the solution. In turn, a
competitive inhibition pattern, or rather a pseudo-competitive inhibition
pattern may result, because the ability of the cellulases to be involved in
the catalysis may be limited due to the lack of available cellulose surface.
In addition the hydrolysis rate may be decreased because of so-called
unproductive, competitive enzyme–substrate binding (Medve et al.,
1998; Ryu et al., 1984). The latter phenomenon, which is also known
from multi-enzymatic degradation of other plant cell wall substrates,
notably pectin (Bagger-Jørgensen and Meyer, 2004; Norsker et al.,
1999), occurs when enzymes adsorb non-productively to the substrate
surface and thus not only become (at least temporarily) inactive, but
also prevent the access of the active enzymes to the substrate. In this
case the addition of substrate will result in the formation of more ES and
more product formation — as seen with ordinary competitive inhibition.
However, this effect would not be a result of successful competition with
the inhibitor but rather due to the supply of new available surface in the
freshly added substrate. At high E/S ratios or at just low substrate
concentrations the cellulase adsorption should not be affected by the
presence of e.g. cellobiose (Zhao et al., 2004). At low E/S ratios, however,
and in turn with relatively high glucose or cellobiose levels – which may
have been added in inhibition studies – inhibition of cellulase
adsorption may arise because of the high glucose and cellobiose levels.
This effect might be considered a genuine competitive inhibition.
6.5. Substrate inhibition and transglycosylation
Transglycosylation and substrate inhibition, the latter including
the unproductive adsorption of enzymes, are in fact not “true”
inhibition mechanisms. However they are both very important
phenomena that decrease the cellulolytic hydrolysis rate and interfere
with the study of inhibition. In practice, the cellobiose substrate
inhibition may be eliminated by maintaining the cellobiose at a
sufficiently low concentration — e.g. by the presence or addition of βglucosidases. The cellulose substrate inhibition is more complex,
however, involving both unproductive adsorption to cellulose as well
as to lignin, and for which no mathematical description currently
exists to our knowledge. The substrate inhibition becomes important
at relatively higher levels of cellulose/cellobiose e.g. at 2–6% (w/v) for
pure cellulose (Huang and Penner, 1991) and >3–10 g/L for cellobiose
(Grous et al., 1985; Hong et al., 1981; Oh et al., 2000). In contrast,
transglycosylation reaction dominates at low enzyme-to-inhibitor
ratios, e.g. at 0.01 gram of active enzyme per g of added glucose
(Andrić et al., 2010) and possibly at high product concentrations.
Transglycosylation can be avoided by performing the inhibition study
with immediate removal of the formed product.
7. Unique features of cellulase action and inhibition
The majority of inhibition studies with cellulases have examined
the events during the initial phase of the overall enzymatic hydrolysis,
i.e. at low extents of conversion, and attempted to assess initial rates
(Table 5). At these conditions only a relatively small number of
productive insoluble cellulose–cellulase ES complex entities may have
formed at the time at which the inhibition of these was assessed. In
turn, relatively high EI (i.e. high EP) levels may result. Consequently, it
may seem as if the cellulolytic enzymes are equally inhibited
irrespective of the substrate presence and the addition of more
(insoluble) cellulose substrate will not relieve the inhibition as it is
seen for competitive inhibition on soluble substrates. The “recorded”
inhibition pattern may therefore be interpreted as non-competitive,
and may even be interpreted to signify the existence of a remote
inhibitor binding site on the enzyme, even if this conclusion is most
likely incorrect. This influence of the substrate nature on the inhibition pattern is supported by results obtained with soluble
“cellulose” substrates, whose solubility, and thus diffusivity, has
been artificially increased, e.g. carboxy-ethyl-cellulose and carboxymethyl-cellulose (Fujii and Shimizu, 1986; Holtzapple et al., 1990). In
these cases, the resulting cellulase inhibition pattern was found to be
competitive (Fujii and Shimizu, 1986; Holtzapple et al., 1990).
7.1. Cellulase binding domain (CBD) and inhibition
All T. reesei cellulases, except EG III, but including most other
known cellulases from other microorganisms, consist of a two domain
structure encompassing a catalytic domain (CD) and a cellulose
binding domain (CBD); the CD and CBD are bound together by a
flexible linker peptide (Linder and Teeri, 1997; Palonen et al., 2004).
P. Andrić et al. / Biotechnology Advances 28 (2010) 308–324
The CBD functions to improve the binding of the CD and in turn
facilitate the activity of the CD on the insoluble cellulose
substrate (Linder and Teeri, 1997). Separate CDs, which have
been identified in culture filtrates of T. reesei (Suurnäkki et al.,
2000) can still hydrolyse the cellulose substrate, however (Palonen
et al., 2004).
The enzyme–substrate complex, either productive (ES) or nonproductive (ES′) (Table 2), can be formed only after the adsorption of
the soluble cellulase (E) onto the insoluble cellulose has taken place.
The adsorbed cellulase, which has not yet reacted, is designated as
[E‐‐‐S] in the discussion below.
When developing their HCH-1 model of enzymatic cellulose
hydrolysis and inhibition, Holtzapple et al. (1984a) used a model
which included enzyme adsorption [E‐‐‐S] and enzyme–substrate
complex formation (ES) as two distinct and separate steps in
cellulose hydrolysis. To describe the adsorption of the free enzyme
onto the substrate surface, Holtzapple et al. (1984a) used the
association (adsorption) constant δ, in which Sf designates the free
cellulose site:
δ=
½E½Sf ½E‐‐S
The model additionally included the possibility of inhibition of all
enzyme species; i.e. the HCH-1 mechanism, and was outlined as:
E + Sf ⇌E‐‐‐S
E‐‐‐S + S⇌ES→E + P
E + P⇌EP
E‐‐‐S + P⇌E‐‐‐SP
ES + P⇌ESP
According to these authors, this inhibition thus represents a noncompetitive type. According to conventional understanding the
competitive inhibitors do not bind to the ES specie. Hence, the
competitive inhibition with the HCH-1 mechanism therefore includes
only the four first equations in the outline of the reactions above
(Holtzapple et al., 1990).
Asenjo (1983) has also presented a model which distinguishes
between the adsorbed, but not yet reacted enzyme, i.e. [E‐‐‐S], and the
true enzyme–substrate complex, ES. In this model, both the soluble
(E) and the adsorbed cellulases ([E‐‐‐S]) were envisaged to be
competitively inhibited by cellobiose and glucose while the productive enzyme–substrate complex (ES) was not subjected to inhibition
by any of the products:
E ⇌E‐‐‐S
E‐‐‐S + S⇌ES→E + P
E + P ⇌EP
E‐‐‐S + P ⇌E‐‐‐SP
It is generally assumed that the CBD directed cellulase adsorption
onto the substrate surface occurs relatively fast, i.e. in the early stages
of hydrolysis (Linder and Teeri, 1997). Many authors have therefore
interpreted cellulase inhibition to be uncompetitive (Tables 3 and 4).
However, rather than a true reaction between the ES and the inhibitor,
the reaction may in reality involve binding of the inhibitory product to
the adsorbed enzyme ([E‐‐‐S]) (Table 7). Essentially, this is therefore a
“competitive inhibition of the adsorbed enzyme” (Table 7) as similar to
the “competitive inhibition of the free enzyme E” (Table 1). Thus, the
terms “uncompetitive” and “competitive” may in this case describe the
same mechanism, adding to the already existing terms ambiguity.
A conceivable explanation of the enzyme adsorption and inhibition is that the adsorbed enzymes are primarily responsible for
cellulose hydrolysis and in turn that the inhibition of the cellulases by
cellobiose and glucose occurs only after the protein adsorption step
(Lee and Fan, 1983) (Table 7). This comprehension builds on that the
319
enzyme molecules become tightly bound to the cellulose particles
immediately upon their contact with cellulose:
E + S⇌ES→E + P
ES + P⇌ESP
ES⇔E‐‐‐S
This assumption implies that the extent of soluble enzyme
adsorption – a measurable quantity – is proportional to the ES complex
formation. The finding that the presence of glucose and/or cellobiose, e.g.
if added during the inhibition study, does not influence the adsorption of
soluble enzyme protein whereas the hydrolysis rates are significantly
reduced by cellobiose and glucose corroborate this theory (see Table 5).
Wald et al. (1984) showed that the cellulase adsorption process is rapid
at the start of hydrolysis, when the product concentration is low, and
essentially proposed the same adsorption and hence inhibition
mechanism, i.e. that [E‐‐‐S] = [ES], as Lee and Fan (1983).
Wald et al. (1984) assumed, however, that the inhibition of
enzyme(s) adsorbed onto the crystalline cellulose fraction was
negligible. On the contrary, Gusakov et al. (1985) argued that the
cellulases display higher susceptibility to the product inhibition when
acting on crystalline cellulose – a result that may indirectly be due to
the lower surface substrate concentration (or specific area) available
for hydrolysis – as compared to amorphous substrates. Similarly,
Johnson et al. (1982) concluded that the effect of cellobiose inhibition
will depend on the nature of the substrate. Bacterial cellulases from
C. thermocellum were for example more inhibited when acting on
microcrystalline than on swollen Avicel.
Gusakov et al. (1992) acknowledged that the sites for adsorption
and catalysis, and hence inhibition, of cellulases are not the same, and
proposed that only the adsorbed cellulase can be inhibited by
cellobiose. In their model, however, they did not distinguish between
the enzyme adsorption and complex formation steps, i.e. [E‐‐‐S] = [ES]
(Gusakov and Sinitsyn, 1992). The same authors did recognize that
the pattern might depend on the adsorption tendency of cellulases
and introduced the dissociation constant Kads, to quantify the cellulose
adsorption ability of cellulases (Gusakov and Sinitsyn, 1992):
Kads =
½E½S
½ES
Cellulases with low Kads (good adsorption ability) can exhibit
uncompetitive inhibition, while cellulases having high Kads (poor
adsorption ability) may exhibit a non-competitive inhibition pattern
(depending also on the E/S ratio, see below).
It is important to note that hardly any of the above authors used
the classical Michaelis–Menten derived methods to determine the
inhibition constants, e.g. via Lineweaver–Burk or Foster–Niemann
plots. Rather, the inhibition constants and the mechanistic interpretation of the events were obtained through kinetic model-fitting of
experimental data over the whole course of hydrolysis, i.e. >8 h of
reaction (Table 5). Apart from product inhibition the models
employed included at least enzyme adsorption and deactivation,
and/or substrate recalcitrance features (Gusakov et al., 1992; Lee and
Fan, 1983; Wald et al., 1984).
7.2. Cellulase catalytic domain (CD) and inhibition
The information about the structure of the catalytic site of cellulases
unambiguously shows that the products can be accommodated and
bind to inhibit the enzyme (Zhao et al., 2004). Furthermore, the tunnel
shaped catalytic site of cellobiohydrolases incorporates multiple sites
for substrate and product binding, that seems to allow for binding of the
product, cellobiose, to the enzyme–substrate complex (productively or
non-productively) or to the enzyme–product complex, even if a remote
site does not exist (Linder and Teeri, 1997).
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P. Andrić et al. / Biotechnology Advances 28 (2010) 308–324
Table 7
The special case of cellulose. Overview of possible product inhibition pathways during
enzyme adsorption and catalysis in cellulase degradation of cellulose. E‐‐‐S—enzyme
adsorbed on cellulose, non-complexed; ES′—non-productive enzyme–substrate complex; ES—productive enzyme–substrate complex (reaction shown in Table 2); E—free
(soluble) enzyme. Except for EP (soluble enzyme–product complex), all other enzyme–
substrate (ES′, ES) and enzyme–substrate-product complexes (E‐‐‐SP, ESP and ES′P),
assume enzyme adsorption prior to complex formation (Table 2).
Possible inhibition interactions
(1) E + P ⇌ EPa
(2) ES + P ⇌ ESPa
(3) ES′ + P ⇌ ES′Pa
(4) EP + S ⇌ ESPa,b
(5) E‐‐‐S + P ⇌ E‐‐‐SPa
(6) E‐‐‐SP ⇌ ESPa,b
a
The combination of each of the enzyme–substrate–product complexes could in
theory bind another P molecule.
b
Theoretically possible.
Gruno et al. (2004) discussed the cellulase inhibition by cellobiose
in the light of the “new” structural information about cellulases
gathered during the 1990s, i.e. mainly in relation to the CBDs and CDs
and the differentiated protein structures, including the multiple
glucosyl-binding tunnel. The structural protein data allow for the
possible existence of a non-productive ES complex — termed as ES′,
even if the cellulase chain is captured by the active site (Gruno et al.,
2004). Assuming that the binding constants for the substrate (for the
forward reaction) are not affected by the presence of the inhibitor –
and vice versa – a mechanism for cellobiohydrolase action on cellulose
was proposed (Gruno et al., 2004). This inhibition model assumes that
the enzyme, irrespective of the enzyme species, can combine to the
cellulose substrate both productively, i.e. immediately correctly
positioned for cleavage, and non-productively, where the substrate
or the enzyme have to move to the correct position:
0
E + S⇌ES ⇌ES→E + P
It is in turn assumed that the reaction can be inhibited by
cellobiose at each of these enzymatic steps (except the ES specie):
E + P⇌EP
EP + S⇌ESP
0
ES + P⇌ESP
Fundamentally, the inhibition then resembles the mixed type of
inhibition (Table 1): If the prevalent productive complex without
inhibitor is productive, a competitive type of inhibition will occur; if it
is non-productive then mixed inhibition or a special case of noncompetitive inhibition will be observed (Gruno et al., 2004). The
dimensionless parameter KES can be used to include an account of the
tendency towards formation of productive enzyme–substrate complex formation:
KES =
½ES0 ½ES
Low KES designates a strong tendency towards productive complex
formation and vice versa. The KES may to a significant extent resemble
the above described enzyme complexing tendency (η) (Holtzapple
et al., 1984a).
Bezerra and Dias (2004) proposed a mechanism for cellobiose
inhibition of cellobiohydrolases similar to the general mixed type of
inhibition kinetics. This mechanism included the existence of a nonproductive ES′ complex reflecting two distinct enzyme–substrate
interactions. They also included the possible existence of an enzyme–
product–product complex (EPP) based on the possibility that
minimum two cellobiose molecules may be bound to the catalytic
domain. This gives a parabolic type of inhibition (Table 1). When
considering the possible existence of a [E‐‐‐S] stage, the product
inhibition could then start via several different pathways, see reactions
e.g. 4), 5), and 6) (Table 7). A detailed analysis of the proposed
parabolic inhibition model has however shown that the parabolic
inhibition and the formation of non-productive complexes were not
the principle constraints limiting the cellulose hydrolysis. The authors
that originally proposed the parabolic mechanism therefore subsequently omitted this mechanism in their subsequent work (Bezerra
and Dias, 2005).
Some of the ambiguity with respect to mechanism of cellulase
inhibition may be avoided if it is accepted that the enzyme can be
inhibited non-competitively or via a mixed type inhibition, while the
catalytic domain is bound to the substrate, i.e. in the [E‐‐‐S] form
(Tables 2 and 6).
As already discussed above, cellulose products must competitively
inhibit the enzymes that catalyze the reaction, as the product structure
is in principle an analogue of the substrate — because it is a “building
block” of the substrate. It is thus expected that this intimate interaction
would involve a product attachment to the same (active) site from
which it was desorbed (Frieden and Walter, 1963). The enzyme product
inhibition should thus theoretically always be of a ‘competitive’ nature
because the product is able to bind the active site of the enzyme —
irrespective of whether this site is activated as in the productive ES
complex. However, even though some products compete for the active
site of the enzyme, they may not necessarily only appear as competitive
inhibitors (Frieden and Walter, 1963). This is why, the reported types of
product inhibition, i.e. non-competitive/mixed, uncompetitive, are
typically related to the existence of remote (control) sites on the
enzymes even though it may be questioned if they really exist.
If the classical definitions can allow that e.g. non-competitive or
mixed type of inhibition may account for product interactions with the
active site, then there would be no conceptual limitations in
understanding that the free cellulase and the cellulase–cellulose
complexes can be inhibited by the product that binds to the active site.
The kinetic description of inhibition will in any case not be affected by
this assumption, as it is not important for the mathematical modeling
whether the inhibitor binds to the active or a remote site. The noncompetitive/mixed type of inhibition could thus be justified to be used
for modeling purposes as it realistically represents the occurrence of
mechanistic enzyme inhibition events and additionally gives better
modeling results (see Section 5.5). Hence, for cellulases, the competitive and uncompetitive product inhibition schemes should be
considered as subtypes of a more generally valid non-competitive/
mixed type of inhibition.
7.3. Unique parameters affecting cellulase inhibition
As shown above, the cellulase inhibition pattern resulting from the
cellobiose and glucose addition studies has only in some cases been
interpreted as competitive (Tables 3 and 4). Rather, it has for a very
long time been proposed to be the uncompetitive and mixed/noncompetitive type that cause an exaggerated quantitative effect on
enzyme reactions (Frieden and Walter, 1963). A major complication
of cellulase inhibition mechanisms is obviously the diverse conclusions that have resulted from the studies discussing the inhibitory
interactions in relation to the reactions between the enzymes and the
substrate (Fig. 4). Perhaps, it is the limited options for experimentally
distinguishing between the interaction of the inhibitor with the free
enzyme, the adsorbed, but not yet reacting enzyme, or with the
enzyme–substrate complex, respectively.
The tendency of the cellulases to be subject to inhibition is
described by different parameters. We have considered the following:
(a) cellulase adsorption ability (Kads) (Gusakov and Sinitsyn, 1992) or
δ (Holtzapple et al., 1984a); (b) cellulose complexing tendency (η)
including cellulose substrate diffusivity (D) (Holtzapple et al., 1990),
see Fig. 3), and (c) cellulose tendency towards productive complex
formation (KES) (Gruno et al., 2004). Thus, it appears that the reported
inhibition might turn out to be competitive if the investigated cellulose
P. Andrić et al. / Biotechnology Advances 28 (2010) 308–324
321
Fig. 4. The summarized overview of conclusions recording cellulose product inhibition patterns drawn from the studies dealing with cellulase inhibition mechanisms in relation to
the action of mainly cellobiohydrolases on cellulosic substrates (Gruno et al., 2004; Holtzapple et al., 1984b, 1990; Gusakov and Sinitsyn, 1992). Product inhibition at RS: remote site
or AS: active site; D: cellulose diffusion coefficient; E/S: enzyme–substrate ratio; Kads: equilibrium constant for adsorption; KES: ratio of non-productive to productive complex. The
part of the scheme that is based on E/S ratio is made according to: high (≈ 1 (w/w)) (Holtzapple et al., 1990), vs. low (≈ 0.01–0.05 (w/w)) value (Gruno et al., 2004).
has higher diffusivity (high D or low η) or if it is forming a productive
ES complex more easily than it is forming an ES′ complex (low KES)
and/or if cellulases are readily adsorbed (low Kads or δ). A competitive
type of inhibition will also be more likely if the E/S ratios are high, in
which case a “pseudo-competitive” inhibition type may occur. Only if
the E/S ratio was low and the enzymes had low Kads (Gusakov et al.,
1992) the observed pattern has been interpreted as uncompetitive. As
we have seen, however, this uncompetitive inhibition essentially
represents “competitive inhibition of adsorbed enzymes”.
Therefore – since competitive inhibition of cellulases seems to
appear with a high substrate solubility or with an increased cellulase
adsorption – the results of cellobiose/glucose inhibition studies could
additionally be used to test e.g. the efficiency of a given pre-treatment
method or the ability of newly developed enzymes to adsorb onto the
substrate, apart from the main purpose of screening product
inhibition of cellulases. Clearly, more experimental work is needed
with this respect, as Fig. 4 is based on a few publications only, which
did not discuss the (broader) extrapolation of the results of inhibition
studies to assess other aspects of cellulase reaction.
7.4. Consequences for reactor design
Contemplation of the available information then leads to an
extended view of the possible inhibitor interactions with the cellulolytic
enzymes as sketched in Fig. 5. Apart from the main reaction pathway
which yields the product (Fig. 5, a1, a2 and a3), the inhibiting product
can thus either bind to the soluble enzyme (E, a1) to give the EP (b1) or
even inhibit/block the very adsorption of the enzyme to the substrate at
high product concentrations (d). It is very likely however that under
these conditions, the active site of the enzyme would also bind the
product (not shown on Fig. 5d). The adsorbed cellulase (E‐‐‐S, a2) might
also be inhibited by the product (E‐‐‐SP, b2) as well as ES (a3) that
proceeds to ESP (b3). The EP complex can on the other hand adsorb onto
the cellulose (b2) and further form a complex, while the product is still
bound to the active site (ESP, b3). The latter interaction does not seem
very likely, however, as the product in this case may block the
productive ES complex formation. It can also be envisaged that the
presence of the product might allow the formation of an ESP complex
but not the release of the product (Fig. 5). On the other hand, if the
product could indeed be formed from ESP the inhibition would resemble
the general hyperbolic type (Table 1). The adsorbed enzyme ([E‐‐‐S])
could also give the non-productive complex (ES′, c1) which might
further be inhibited by the product (ES′P, c2). Finally, all product
complexes with enzyme and substrate (EP, E‐‐‐SP, ESP and ES′P) could
further accommodate at least one extra molecule of the inhibiting
product (not shown in Fig. 5).
Both glucose and cellobiose may apparently bind to the active site
of the cellulase, in almost all complexed forms (Fig. 5), and severely
‘non'competitively inhibit the catalytic action (Table 5). Consequently, the inevitable strategy to diminish the inhibition and increase
the hydrolysis rates is to remove these products from the reaction
environment. Various reactor and reaction set-ups have been
evaluated for accomplishing product removal during enzyme-catalyzed cellulosic hydrolysis reactions - as reviewed in detail in a
separate review (Andrić et al., submitted for publication). Although a
number of issues beyond product inhibition, such as overcoming mass
transfer limitations at high viscosities, become important for reactor
design at high-solids loadings, fast and complete product removal is
particularly important where a high glucose yield is essential for
ethanol production (Fig. 1). The cellobiose concentration may be
minimized by addition of sufficient β-glucosidase activity to convert it
to glucose, but the glucose must then be removed by e.g. application of
membrane reactors to maintain a high conversion rate, high yields,
and high volumetric productivity of the reactor, i.e. relatively high
kgglucose/(m3reactor volume h) that can be obtained with such recalcitrant
substrate.
Even though the cellulase inhibition might be competitive in
nature, the addition of higher levels of substrate to overcome it, is thus
not an appropriate approach because the higher substrate concentrations inherently result in higher product concentrations and hence
Fig. 5. Schematic representation of possible scenarios for product inhibition of cellulases, when the inhibitor is initially present in the mixture. (E)—free (soluble) enzyme, (E‐‐‐S)—adsorbed enzyme, (ES)—enzyme–substrate productive
complex, (ES′)—enzyme–substrate unproductive complex, (EP)—enzyme–product complex, (E‐‐‐SP)—adsorbed enzyme–product complex, (ESP, ES′P)—enzyme–substrate–product complex.
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higher inhibition! Thus, with respect to product inhibition of any type,
the removal of the product inhibitor from the reacting system is the
best alternative for minimization of inhibition.
systems for enzymatic lignocellulose hydrolysis, it might prove fruitful
to attempt to identify or develop new cellulolytic enzymes that are more
resistant to product inhibition than the ones currently used.
8. Minimizing cellulase inhibition: Conclusions
References
One of the main suggestions that for some time has been
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in lignocellulose conversion processes has been that it is crucial to
conduct the enzymatic saccharification at high-solids concentrations
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significant quantitative significance of the product inhibition inherently limits the bioreactor goals of this approach, particularly in batch
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