Development of Highly Sensitive Fluorescent Sensors for Separation-Free Detection and Quantitation Systems of Pepsin Enzyme Applying a Structure-Guided Approach

Abstract

Two fluorescent molecularly imprinted polymers (MIPs) were developed for pepsin enzyme utilising fluorescein and rhodamine b. The main difference between both dyes is the presence of two (diethylamino) groups in the structure of rhodamine b. Consequently, we wanted to investigate the effect of these functional groups on the selectivity and sensitivity of the resulting MIPs. Therefore, two silica-based MIPs for pepsin enzyme were developed using 3-aminopropyltriethoxysilane as a functional monomer and tetraethyl orthosilicate as a crosslinker to achieve a one-pot synthesis. Results of our study revealed that rhodamine b dyed MIPs (RMIPs) showed stronger binding, indicated by a higher binding capacity value of 256 mg g −1 compared to 217 mg g −1 for fluorescein dyed MIPs (FMIPs). Moreover, RMIPs showed superior sensitivity in the detection and quantitation of pepsin with a linear range from 0.28 to 42.85 µmol L −1 and a limit of detection (LOD) as low as 0.11 µmol L −1. In contrast, FMIPs covered a narrower range from 0.71 to 35.71 µmol L −1 , and the LOD value reached 0.34 µmol L −1 , which is three times less sensitive than RMIPs. Finally, the developed FMIPs and RMIPs were applied to a separation-free quantification system for pepsin in saliva samples without interference from any cross-reactors.

Full text

Citation: Mostafa, A.M.; Barton, S.J.;
Wren, S.P.; Barker, J. Development of
Highly Sensitive Fluorescent Sensors
for Separation-Free Detection and
Quantitation Systems of Pepsin
Enzyme Applying a Structure-Guided
Approach. Biosensors 2024,14, 151.
https://doi.org/10.3390/bios14030151
Received: 6 March 2024
Revised: 13 March 2024
Accepted: 18 March 2024
Published: 20 March 2024
Copyright: © 2024 by the authors.
Licensee MDPI, Basel, Switzerland.
This article is an open access article
distributed under the terms and
conditions of the Creative Commons
Attribution (CC BY) license (https://
creativecommons.org/licenses/by/
4.0/).
biosensors
Article
Development of Highly Sensitive Fluorescent Sensors for
Separation-Free Detection and Quantitation Systems of Pepsin
Enzyme Applying a Structure-Guided Approach
Aya M. Mostafa 1,2,* , Stephen J. Barton 1, Stephen P. Wren 1and James Barker 1
1School of Life Sciences, Pharmacy and Chemistry, Kingston University, Kingston upon Thames,
London KT1 2EE, UK; s.barton@kingston.ac.uk (S.J.B.); s.wren@kingston.ac.uk (S.P.W.);
j.barker@kingston.ac.uk (J.B.)
2Department of Pharmaceutical Analytical Chemistry, Faculty of Pharmacy, Assiut University,
Assiut 71526, Egypt
*Correspondence: ph.ayamoneer@gmail.com
Abstract: Two fluorescent molecularly imprinted polymers (MIPs) were developed for pepsin enzyme
utilising fluorescein and rhodamine b. The main difference between both dyes is the presence of two
(diethylamino) groups in the structure of rhodamine b. Consequently, we wanted to investigate the
effect of these functional groups on the selectivity and sensitivity of the resulting MIPs. Therefore,
two silica-based MIPs for pepsin enzyme were developed using 3-aminopropyltriethoxysilane as
a functional monomer and tetraethyl orthosilicate as a crosslinker to achieve a one-pot synthesis.
Results of our study revealed that rhodamine b dyed MIPs (RMIPs) showed stronger binding,
indicated by a higher binding capacity value of 256 mg g
−1
compared to 217 mg g
−1
for fluorescein
dyed MIPs (FMIPs). Moreover, RMIPs showed superior sensitivity in the detection and quantitation
of pepsin with a linear range from 0.28 to 42.85
µ
mol L
−1
and a limit of detection (LOD) as low
as 0.11
µ
mol L
−1
. In contrast, FMIPs covered a narrower range from 0.71 to 35.71
µ
mol L
−1
, and
the LOD value reached 0.34
µ
mol L
−1
, which is three times less sensitive than RMIPs. Finally, the
developed FMIPs and RMIPs were applied to a separation-free quantification system for pepsin in
saliva samples without interference from any cross-reactors.
Keywords: fluorescent molecularly imprinted polymers; fluorescent biosensors; spectrofluorometric
analysis; pepsin; biomarkers; gastroesophageal reflux disease (GERD)
1. Introduction
Molecular imprinting technology is an innovative technique that has provided syn-
thetic and highly selective receptors for different target analytes. Molecular imprinting is
a simple technique that depends on the copolymerisation of functional monomer(s) and
crosslinker(s) in the presence of the target analyte as a template. Herein, non-covalent
bonds are mainly established between the template and the functional monomer that are
the bases for molecular identification [
1
]. Upon polymerisation, complementary binding
sites are created for the target analyte in the structure of the polymer, which, after removing
the template using a suitable solvent, become available for binding this analyte in different
samples [
2
–
5
]. The growing interest in molecular imprinting technology is due to the
increased advantages it offers in comparison to their biological alternative, antibodies. Up
till the present, antibodies have been used for the detection of different biological analytes,
specifically through enzyme-linked immunosorbent assay (ELISA), due to their extremely
high selectivity. However, the sensitivity of antibodies to heat and pH, their high cost of
production, and their short shelf life are significant drawbacks. Moreover, some antibodies
need to be labelled for the detection of the binding event, which is not a straightforward
Biosensors 2024,14, 151. https://doi.org/10.3390/bios14030151 https://www.mdpi.com/journal/biosensors

Biosensors 2024,14, 151 2 of 18
procedure and may result in an alteration of the binding properties of the antibodies. There-
fore, molecular imprinting technology proved to be an ideal alternative for antibodies as
molecularly imprinted polymers (MIPs) are more stable, cost-effective, more resistant to
degradation, and can be tailor-made to bind any analyte of interest [6].
Labelling of MIPs with a fluorescent reporter can be a very useful tool that helps to
translate the binding event between MIPs and the target into a readable signal. Creating
fluorescent MIPs can be achieved through three major approaches: (1) incorporating fluo-
rescent particles into the core of the developed polymer, such as carbon dots [
7
,
8
], quantum
dots [
9
–
12
], or gold nanoparticles [
13
] in a technique known as core-shell imprinting; (2) the
use of a fluorescent functional monomer in the polymerisation mixture [
14
–
16
]; or (3) post
imprinting modification of the resulting MIPs with a fluorescent dye [
17
]. Nonetheless, the
first two approaches are more popular and involve less risk of alteration of the binding
properties of the developed MIPs [
18
]. Fluorescent MIPs not only allow readable detection
of the binding event but also the exploitation of the fluorescence as a quantitation method
for the concentration of the target. In this case, the binding of the target to fluorescent MIPs
results in an alteration of the fluorescence, either an increase (less common) or a decrease in
the fluorescence intensity, which is usually proportional to the concentration of the analyte.
Therefore, fluorescent MIPs provide not only a method for extraction and detection of the
target analyte but also a very sensitive and accurate method of quantitation, which makes
them a very inclusive and self-sufficient technique of analysis [19].
Fluorescein is a widely known fluorescent dye that belongs to the xanthene class
of dyes, which has been used for decades in fluorescent labelling. The incorporation of
fluorescein into the structure of MIPs has been successfully reported multiple times to
determine different targets, owing to its strong fluorescence and market availability [
20
]. In
the paper reported by F. Wang et al., fluorescein has been applied in the development of
fluorescent MIPs for the detection of naproxen, a common non-steroidal anti-inflammatory
drug, through a simple and catalyst-free sol-gel polymerisation method [
21
]. The authors
acknowledged that the achieved limit of detection for the developed MIPs was not satis-
factory in comparison to other methods reported for naproxen, and they suggested the
use of a more sensitive dye. Therefore, in light of this recommendation, we present the
development of two fluorescent MIPs utilising fluorescein and rhodamine b as fluorescent
dyes for the detection of the pepsin enzyme. Fluorescein and rhodamine b are conjugated
through a simple reaction with (3-aminopropyl)trimethoxysilane to create the fluorescent
co-monomer. This fluorescent co-monomer is incorporated in a one-pot sol-gel polymerisa-
tion method to produce the yellow-coloured FMIPs from fluorescein and the pink-coloured
RMIPs from rhodamine b.
Pepsin was chosen as a target for our work because it has been recently confirmed
as a biomarker for gastroesophageal reflux disease (GERD). GERD is a common disorder
occurring in over 20% of the world’s population [
22
]. The early diagnosis and treatment
of GERD can efficiently contribute to the prognosis of this common disease. The pepsin
enzyme is one of the earliest discovered enzymes that aid in the digestion of proteins in
humans and animals. However, the enzyme can also cause increased damage to the mucosal
lining of the stomach in patients with GERD, resulting in more pain [
23
]. Furthermore, by
analysing the saliva of patients with GERD, it was found that the saliva exhibited higher
levels of pepsin enzyme compared to healthy individuals. As a result, elevated levels of
pepsin in the saliva of patients could be used as a biomarker for GERD and act as an early,
simple, and non-invasive diagnostic tool [
24
]. Up till now, the main method of measuring
the level of pepsin and diagnosis of GERD is the Peptest
®
, which is an ELISA method
utilising antibodies [
25
]. However, we aim to find a synthetic, cheaper, and more stable
alternative to conventional antibody testing.
Our group has recently focused on the synthesis of MIPs as extraction and/or analyti-
cal tools for pepsin as an important biomarker for GERD. Imprinting of proteins can be
challenging due to their big size, pH sensitivity, conformational stability, and limited solu-
bility [
26
]. Nonetheless, there has been a huge progress in the field of imprinting of proteins

Biosensors 2024,14, 151 3 of 18
through the use of more targeted monomers, suitable solvents and epitope imprinting [
27
].
In our review of the literature, very few papers dealt with the imprinting of pepsin [
28
,
29
],
of which only one paper described the development of fluorescent MIPs for the detection
of pepsin in an ELISA-like format [
30
]. In this reported study, the analysis of pepsin was
based on the binding of fluorescent pepsin-specific MIPs to magnetic pepsin nanoparticles
immobilised on magnetic inserts in the wells of a microtiter plate. Herein, free pepsin in the
sample competed with immobilised pepsin, causing a decrease in fluorescent MIPs binding
to the magnetic inserts, hence increasing the central fluorescence of the well. The synthesis
of fluorescent MIPs in this reported work was not straightforward and included multiple
steps. There was no optimisation of the concentration or types of monomers or crosslinkers
and no justification for the choice of monomers applied in their method. In addition, no
chemical, thermal, or functional characterisation profiles, including kinetics and isotherm,
were provided. Finally, there was no application on human saliva samples. In one of our
previous works, we developed magnetic MIPs for the extraction of pepsin, utilising the
core-shell imprinting technique [
31
]. The developed magnetic MIP bound pepsin in saliva
samples, which was then released from MIPs and analysed via high-performance liquid
chromatography. However, we aspired to design fluorescent MIPs that would be capable of
both extraction and analysis of pepsin in a single step. Therefore, in this current work, we
developed two fluorescent MIPs using fluorescein and rhodamine b as fluorescent reporters
with complete optimisation and characterisation profiles. In addition, analytical method
optimisation and validation were investigated for the determination of pepsin using both
MIPs to compare their analytical performance. Finally, the fluorescent MIPs were used
to demonstrate their applicability for the extraction and quantitation of pepsin in human
saliva samples.
2. Experimental Section
2.1. Materials and Instrumentation
Tetraethyl orthosilicate for synthesis (TEOS), sodium dodecyl sulphate 99% (SDS),
(3-aminopropyl)triethoxysilane 98% (APTES), deionised water, absolute ethanol, sodium
chloride 99.5%, phosphate-buffered saline (PBS) tablets (pH 7.2), human pepsin, human
lipase, and human amylase were all procured from Thermo Fisher Scientific (Horsham,
UK). Fluorescein-5-isothiocyanate 98% (FITC) and rhodamine b isothiocyanate 98% (RITC)
were purchased from Fluorochem, (Hadfield, Glossop, UK).
Fluorescence measurements were carried out using a Cary Eclipse fluorescence spec-
trometer (Agilent, London, UK), running on a Cary Eclipse software version 1.1 (132)
(Agilent, London, UK). UV measurements were conducted using a Cary UV-Vis Compact,
operating on a Cary UV Workstation™, software version 1.0.1284 (Agilent, London, UK).
Thermal characterisation, including thermogravimetric experiments, was performed on
a Mettler Toledo TGA/DSC 1 Series (Leicester, UK)—running on STARe™ software Ver-
sion 10.00 (Mettler Toledo, Leicester, UK), and differential scanning calorimetric assays
(DSC) were performed on a TA Instruments DSC25 Series (New Castle, UK), running on
Trios™ software, v5.4.0.300 (TA instruments, New Castle, UK) Infra-red analysis was con-
ducted using a Thermo Fisher (Horsham, UK) Scientific Nicolet iS5 Fourier transform
infrared spectroscopy (FTIR) running on OMNIC™ software version 9.13.5.1294 (Ther-
mofisher, Horsham, UK).
13
C Nuclear magnetic resonance (NMR) analysis was carried out
using a Bruker Avance III 600 two-channel FTNMR spectrometer (Coventry, UK) operating
at 600 MHz and utilising TopSpin™ software version 4.3.0 (Bruker, Coventry, UK) for
data analysis. Scanning electron microscopy (SEM) was used to visualise the resulting
polymers using a Zeiss Evo-50 electron microscope (Cambridge, UK) operating on Smart
SEM™ software version 5.0 (Zeiss, Cambridge, UK). Data processing and graph plotting
were executed using Origin™ 8.5 software (Origin Lab Corporation, North Hampton,
NH, USA).

Biosensors 2024,14, 151 4 of 18
2.2. Synthesis of Fluorescent Co-Monomers
Two fluorescent co-monomers were prepared independently from a simple coupling
reaction between APTES and FITC or RITC according to a previously reported method [
32
].
Each dye (0.04 mmol) was dissolved in 8 mL absolute ethanol, and 0.04 mmol of APTES
was added to the solution and continuously stirred for 24 h. The obtained product was
used without further purification.
2.3. Preparation of Fluorescent MIPs for Pepsin
Synthesis of fluorescent MIPs was carried out through a simple sol-gel polymerisation
reaction, which is an effective, simple and green method [
4
]. APTES was employed as a
functional monomer, and TEOS as a crosslinker. The amounts and volumes of different
reagents used in the synthesis procedure were carefully optimised to reach the best possible
binding. The optimised procedure for fluorescein dyed MIPs (FMIPs) was as follows:
15 mg of the target analyte pepsin were dissolved in 8 mL PBS buffer solution (pH 7.2)
followed by the addition of 0.50 mL APTES and stirring for half an hour for preassembly
followed by addition of 9 mL of the fluorescent co-monomer (FITC-APTES) and stirring
for another half hour. Finally, 0.9 mL of TEOS dissolved in 3.5 mL of absolute ethanol was
added dropwise, and the reaction mixture was sealed under a nitrogen atmosphere and
continuously stirred for 48 h. The optimised procedure for rhodamine b dyed MIPs (RMIPs)
was almost the same with different optimum volumes. Pepsin (15 mg) was dissolved in
8 mL PBS buffer (pH 7.2), followed by the addition of 0.75 mL APTES and stirred for half an
hour, followed by the addition of 8 mL of the fluorescent co-monomer (RITC-APTES) and
stirring for another half hour. Finally, 0.9 mL of TEOS dissolved in 3.5 mL absolute ethanol
was added dropwise, and the reaction mixture was sealed under a nitrogen atmosphere and
continuously stirred for 48 h. Fluorescent non-imprinted polymers (NIPs) were prepared
for both FMIPs and RMIPs without the addition of pepsin for comparison purposes. The
resulting MIPs and NIPs were rinsed with deionised water and ethanol twice to remove
remnants of starting materials and oligomers. Furthermore, to remove the target from the
imprinted cavities, both fluorescent MIPs were washed with a solution of 1% w/vsodium
dodecyl sulphate (SDS)/10% v/vacetic acid for 4 h, followed by washing with deionised
water multiple times to eliminate remnants of the washing solution. UV spectrometry
was used to ensure complete washing by testing fragments of the washing solution. The
resulting polymers were dried under vacuum at 60
◦
C and then finely ground and stored
in dark containers to prevent photobleaching.
2.4. Protein Binding Experiments
To assess the successful imprinting, individual binding assays were performed for all
the developed polymers against a known concentration of pepsin. The binding experiments
were carried out by incubating 50 mg of the developed polymers with 20 mL of 1 mg mL
−1
pepsin solution for 2 h (for fluorescein-dyed polymers) and 1 h (for rhodamine b dye
polymers). After shaking for the optimum binding time, solutions were centrifuged at
4500 rpm for 10 min, and the concentration of pepsin in the supernatant was measured by
UV spectrometry against a blank of deionised water incubated with MIPs or NIPs for the
same amount of time. To evaluate the binding capacity of MIPs and NIPs, the amount of
pepsin adsorbed per gram of polymers was calculated using the following equation:
Q=(Ci −Ct)·V/m
where, Q(mg g
−1
) is the quantity of pepsin in milligrams adsorbed per gram of polymer,
Ci (mg mL
−1
) is the starting concentration of pepsin, Ct (mg mL
−1
) is the remaining
concentration of pepsin after incubation time (t), V(mL) is the volume of pepsin solution,
and m(g) is the mass of MIPs or NIPs applied in the experiment. All the experiments were
conducted in triplicate to validate the precision of the results.

Biosensors 2024,14, 151 5 of 18
2.5. Characterisation of the Fluorescent Polymers
Characterisation of the developed fluorescent polymers is a very important step in
comparing the properties of FMIPs and RMIPs, especially the different binding parameters.
Therefore, the morphology of the developed polymers was assessed using SEM imaging,
which also enabled an estimation of the particle size.
13
C nuclear magnetic resonance (NMR)
was conducted on FITC, RITC, FITC-APTES, and RITC-APTES to verify the formation of the
thiourea linkage between FITC or RITC with APTES to form the fluorescent co-monomers.
Fourier transform infrared spectra (4000–500 cm
−1
) were collected for FITC, RITC, FMIPs,
FNIPs, RMIPs, and RNIPs to compare the prominent bands, ensure template removal, and
verify the absence of any residual starting materials.
Thermogravimetric analysis (TGA) and differential scanning calorimetry (DSC) were
performed on FITC, RITC, FMIPs, FNIPs, RMIPs, and RNIPs. Data provided by TGA and
DSC can ensure complete polymerisation, detect any unreacted starting materials, and
determine the content of adsorbed moisture. TGA was performed along a temperature
range from 25 to 650
◦
C at a heating pace of 10
◦
C min
−1
and a nitrogen gas flowing at
a rate of 50 mL min
−1
, and DSC was run at a temperature range from 25 to 350
◦
C at a
heating rate of 10 ◦C min−1.
To determine the order of binding kinetics, rebinding experiments were performed on
the developed polymers at increasing time intervals, I.e., 50 mg of the fluorescent polymers
were incubated with 20 mL of 1 mg mL
−1
pepsin solution for 0, 1, 2, 4, 6, and 8 h. The
quantity of pepsin adsorbed per gram (Q) for each time interval was plotted against time
to determine the binding order kinetics.
Additionally, rebinding experiments were conducted on the developed polymers by
applying different concentrations of pepsin to determine the binding isotherm model. Thus,
50 mg of the fluorescent polymers were incubated with 20 mL of pepsin solutions in the
concentration range (0.2 to 1.5 mg mL
−1
) for 1 h (for RMIPs and RNIPs) and 2 h (for FMIPs
and FNIPs). The quantity of pepsin adsorbed per gram (Q) for each concentration was
plotted against concentration to depict the binding isotherm model.
To verify the binding selectivity of FMIPs and RMIPs, their binding was compared
to the binding of their corresponding FNIPs and RNIPs, respectively. Subsequently, the
imprinting factor (IF) was calculated by dividing the value of Q for MIPs by that of NIPs.
In addition, since the target analyte in this work is salivary pepsin, it is only reasonable
to test the selectivity of the developed fluorescent polymers against other proteins that
can exist with pepsin in human saliva. Amylase and lipase, along with other enzymes,
were selected to test the selectivity due to their abundance in saliva and possible cross-
reactivity. Therefore, a similar binding assay for pepsin was performed on the competitor
enzymes where 20 mL of 1 mg mL
−1
solution of each enzyme was incubated with 50 mg of
the fluorescent polymers for 1 h (for RMIPs and RNIPs) and 2 h (for FMIPs and FNIPs).
The quantity bound per gram (Q) was calculated for each enzyme and compared to that
of pepsin.
2.6. Stability Testing
The developed MIPs were tested for their stability and shelf life to verify their suitabil-
ity for long-term use. The developed FMIPs and RMIPs were left for 0- 1-, 3-, and 6-month
periods after preparation while being stored in dark containers. In addition, other batches
of FMIPs and RMIPs were stored at 10, 25, 35
◦
C and 45
◦
C for 1 month period to test for
stability against storage temperatures. After these storage periods, the binding capacity of
pepsin was tested using the procedure for protein binding assay mentioned previously.
2.7. Fluorescence Measurements
The procedure for measuring the fluorescence intensity of the developed fluorescent
polymers was carefully optimised to get the highest possible sensitivity for FMIPs and
RMIPs. Consequently, a suspension of the fluorescent polymers in water (3 mg mL
−1
)
was prepared, centrifuged to remove coagulated particles, and measured at an excitation

Biosensors 2024,14, 151 6 of 18
wavelength (
λexc
) of 471 nm for FMIPs and 546 nm for RMIPs at PMT voltage of 600 and
slit width of 10 nm for both excitation and emission. In order to construct a calibration
curve, different concentrations of pepsin were added to the suspension of polymers for the
optimum binding time of 2 h or 1 h for FMIPs and RMIPs, respectively, with continuous
shaking. Relative fluorescence intensity was measured for each sample and plotted against
the concentration of pepsin to establish the linear range and deduct the regression equation.
2.8. Application to Measuring Pepsin in Human Saliva
This study received ethical consent from Kingston University Ethics Committee (Ethics
Code 2895) and was conducted in concordance with the regulations of the UK Human
Tissue Act (HTA) 2004. Saliva samples were collected, centrifuged at 4500 rpm for 30 min,
and used immediately. 100
µ
L of saliva were spiked with increasing concentrations of
pepsin in the concentration range (0–42.85
µ
mol L
−1
). One mL of the suspension of FMIPs
or RMIPs (3 mg mL
−1
) was added to each sample and incubated for the optimum binding
time. Each sample was measured in triplicate using a spectrofluorometer to ensure the
precision of the results.
3. Results and Discussion
3.1. Preparation of FMIPs and RMIPs
The use of fluorescein as a fluorescent dye for manufacturing of MIPs was previously
reported for naproxen [
20
]. However, the authors of this paper stated that the use of
fluorescein did not achieve the desired sensitivity levels and that other dyes needed to
be tested. Based on this recommendation, we tested another cheap and readily available
organic dye, rhodamine b, along with fluorescein. Although the structure of rhodamine
b is very close to that of fluorescein, it still has some structural merit that we presumed
can enhance binding and sensitivity. A neutral pH was chosen as the working pH since
we detect pepsin in saliva, which has a neutral pH (6.2–7.6) [
33
]. As pictured in Figure 1,
rhodamine b has two diethylamino groups in its structure, which are positively charged at
the neutral working pH (7.2). Pepsin has a relatively low isoelectric point of 3.24 [
34
], which
means that it is negatively charged at pH 7.2. Therefore, a strong electrostatic interaction
was predicted between rhodamine b and pepsin, unlike fluorescein, which can only interact
with pepsin through hydrogen bonds in its carboxylic group.
Biosensors2024,14,xFORPEERREVIEW7of19
Figure1.Schematicpresentationofpolymerisationandinteractionpointsbetweenpepsinandflu-
oresceinorrhodamineb.
APTESwaschosenasafunctionalmonomerduetoitsabilitytointeractwiththe
aminoandcarboxylicgroupsinthebackboneofthepepsinmoleculeviahydrogenbonds.
Moreover,APTESwasalsocoupledtoFITCorRITCtogeneratethefluorescentco-mon-
omer,whichisalsocapableofhydrogenbondingwiththetarget.Asaresult,afterwash-
ingthedevelopedFMIPsorRMIPsandremovalofthetarget,manybindingsitescomple-
mentarytopepsinwouldbegenerated.Inaddition,fluoresceinorrhodaminebmolecules
wouldbedistributedinthepolymermatrixwithhighprevalenceinthebindingsitesdue
totheprecedenthydrogenand/orelectrostaticbondsformedwiththetargetduringthe
polymerisationphase.Consequently,thebindingandthereleaseofpepsintothebinding
siteswouldresultinasignificantchangeinthefluorescenceintensity,whichcanberec-
ordedandutilisedtodetectthepresenceofpepsinanddetermineitsconcentration.Ad-
ditionally,aslightcolourchangebetweenMIPsandtheircorrespondingNIPswasno-
ticed,inwhichMIPsalwaysseemeddarkerincolourthanNIPs,asshowninthepictures
inFigureS1.Thiscanbeaributedtothegreaterconcentrationofthefluorescentco-mon-
omerinthepolymericmatrixofMIPscomparedtoNIPs;thisobservationliesinconcord-
ancewiththesameobservationsforfluoresceinimprintedpolymerspreviouslyprepared
fornaproxen[20].Sincethepepsinmoleculecanformmultiplebondswiththefluorescent
co-monomer,itispredictedthatthepresenceofpepsinresultedinhigherpolymerisation
efficiencyofFITC-APTESorRITC-APTESintoMIPs,resultinginadarkercolour.
Allthereagentsusedinthesynthesisprocedurewereinvestigatedthoroughlyto
studytheireffectonthebindingcapacity(Q).DifferentvolumesofAPTES,FITC-APTES
orRITC-APTES,andTEOSweretested,aswellasdifferentpreassemblytimesanddiffer-
entamountsofpepsintofullyoptimisetheprocedure.FiguresS2andS3showtheopti-
mumvaluesobtainedfromthedifferentoptimisationexperimentsforbothfluorescein-
dyedpolymersandrhodamineb-dyedpolymers,respectively.Inaddition,testingdiffer-
entwashingsolutionssuchassodiumchloride(0.5molL−1),phosphatebuffer(0.05mol
L−1,pH7.2),andasolutionof1%w/vSDS/10%v/vaceticacidwasnecessarytofindthe
solutionthatremovesalltracesofthetemplate.Thisisverycrucialasincompletetemplate
removalresultsintemplatebleeding,whichcancausesignificanterrorintheanticipated
resultand/orblockageoftheavailablebindingsites,leadingtoareductioninsensitivity.
Therefore,aftermeticuloustestingofthedifferentwashingsolutionsandtimeofwashing,
Figure 1. Schematic presentation of polymerisation and interaction points between pepsin and
fluorescein or rhodamine b.

Biosensors 2024,14, 151 7 of 18
APTES was chosen as a functional monomer due to its ability to interact with the amino
and carboxylic groups in the backbone of the pepsin molecule via hydrogen bonds. More-
over, APTES was also coupled to FITC or RITC to generate the fluorescent co-monomer,
which is also capable of hydrogen bonding with the target. As a result, after washing the
developed FMIPs or RMIPs and removal of the target, many binding sites complementary
to pepsin would be generated. In addition, fluorescein or rhodamine b molecules would be
distributed in the polymer matrix with high prevalence in the binding sites due to the prece-
dent hydrogen and/or electrostatic bonds formed with the target during the polymerisation
phase. Consequently, the binding and the release of pepsin to the binding sites would result
in a significant change in the fluorescence intensity, which can be recorded and utilised
to detect the presence of pepsin and determine its concentration. Additionally, a slight
colour change between MIPs and their corresponding NIPs was noticed, in which MIPs
always seemed darker in colour than NIPs, as shown in the pictures in Figure S1. This can
be attributed to the greater concentration of the fluorescent co-monomer in the polymeric
matrix of MIPs compared to NIPs; this observation lies in concordance with the same
observations for fluorescein imprinted polymers previously prepared for naproxen [
20
].
Since the pepsin molecule can form multiple bonds with the fluorescent co-monomer, it
is predicted that the presence of pepsin resulted in higher polymerisation efficiency of
FITC-APTES or RITC-APTES into MIPs, resulting in a darker colour.
All the reagents used in the synthesis procedure were investigated thoroughly to
study their effect on the binding capacity (Q). Different volumes of APTES, FITC-APTES or
RITC-APTES, and TEOS were tested, as well as different preassembly times and different
amounts of pepsin to fully optimise the procedure. Figures S2 and S3 show the optimum
values obtained from the different optimisation experiments for both fluorescein-dyed
polymers and rhodamine b-dyed polymers, respectively. In addition, testing different
washing solutions such as sodium chloride (0.5 mol L
−1
), phosphate buffer (0.05 mol L
−1
,
pH 7.2), and a solution of 1% w/vSDS/10% v/vacetic acid was necessary to find the
solution that removes all traces of the template. This is very crucial as incomplete template
removal results in template bleeding, which can cause significant error in the anticipated
result and/or blockage of the available binding sites, leading to a reduction in sensitivity.
Therefore, after meticulous testing of the different washing solutions and time of washing,
a solution of 1% w/vSDS/10% v/vacetic acid was determined to be the most effective in
the removal of the template within 4 h.
3.2. Characterisation of the Fluorescent MIPs
3.2.1. Morphological Characterisation
SEM images were collected for FMIPs, FNIPs, RMIPs, and RNIPs and are displayed in
Figure 2. As the images show, the generated polymers are not entirely spherical, with a
rough surface and some scattered, coagulated portions. This is due to the application of
the bulk polymerisation technique in which there is no complete control over the size or
morphology of the synthesised polymers. Moreover, a basic estimation of the particle size
was made using the resulting SEM pictures. Here, we can report that the particles have a
size ranging from 0.5 to 2.0 microns, which again is typical for bulk polymerisation.
To investigate the effect of the variability of particle size on fluorescence signal re-
producibility, a one-way analysis of variance (ANOVA) test was conducted. One batch
(3 mg mL
−1
) of each of the FMIPs, FNIPs, RMIPs, and RNIPs was prepared, and the
fluorescence signal was measured three times for each solution. The same experiment was
conducted every day over a week period. The collected data for seven days (seven groups)
for each polymer type was tested via ANOVA analysis, and the results are displayed in
Table 1. Since the F-statistic values are less than the critical values, they suggest that the
differences between the group means are not statistically significant. This implies that
there is no strong evidence to reject the null hypothesis, and we conclude that there are no
statistically significant differences between the means of the groups for all polymers. In

Biosensors 2024,14, 151 8 of 18
other words, the variation observed between the groups could likely be due to random
chance alone, and there may not be real differences in the means of the groups.
Biosensors2024,14,xFORPEERREVIEW8of19

asolutionof1%w/vSDS/10%v/vaceticacidwasdeterminedtobethemosteffectivein
theremovalofthetemplatewithin4h.
3.2.CharacterisationoftheFluorescentMIPs
3.2.1.MorphologicalCharacterisation
SEMimageswerecollectedforFMIPs,FNIPs,RMIPs,andRNIPsandaredisplayed
inFigure2.Astheimagesshow,thegeneratedpolymersarenotentirelyspherical,with
aroughsurfaceandsomescaered,coagulatedportions.Thisisduetotheapplicationof
thebulkpolymerisationtechniqueinwhichthereisnocompletecontroloverthesizeor
morphologyofthesynthesisedpolymers.Moreover,abasicestimationoftheparticlesize
wasmadeusingtheresultingSEMpictures.Here,wecanreportthattheparticleshavea
sizerangingfrom0.5to2.0microns,whichagainistypicalforbulkpolymerisation.

Figure2.SEMpicturesofFMIPs,FNIPs,RMIPs,andRNIPs.
Toinvestigatetheeffectofthevariabilityofparticlesizeonfluorescencesignalrepro-
ducibility,aone-wayanalysisofvariance(ANOVA)testwasconducted.Onebatch(3mg
mL−1)ofeachoftheFMIPs,FNIPs,RMIPs,andRNIPswasprepared,andthefluorescence
signalwasmeasuredthreetimesforeachsolution.Thesameexperimentwasconducted
everydayoveraweekperiod.Thecollecteddataforsevendays(sevengroups)foreach
polymertypewastestedviaANOVAanalysis,andtheresultsaredisplayedinTab l e 1.
SincetheF-statisticvaluesarelessthanthecriticalvalues,theysuggestthatthedifferences
betweenthegroupmeansarenotstatisticallysignificant.Thisimpliesthatthereisno
strongevidencetorejectthenullhypothesis,andweconcludethattherearenostatisti-
callysignificantdifferencesbetweenthemeansofthegroupsforallpolymers.Inother
words,thevariationobservedbetweenthegroupscouldlikelybeduetorandomchance
alone,andtheremaynotberealdifferencesinthemeansofthegroups.

Figure 2. SEM pictures of FMIPs, FNIPs, RMIPs, and RNIPs.
Table 1. ANOVA parameters for testing the effect of particle size variability on the reproducibility of
the fluorescence signal for FMIPs, FNIPs, RMIPs, and RNIPs.
Polymer SSB SSW SST F Calculated F Critical (α= 0.05)
FMIPs 47.195 180.125 227.321 0.611 2.847
FNIPs 3.619 13.163 16.782 0.6415 2.847
RMIPs 7.272 24.678 31.950 0.687 2.847
RNIPs 6.404 60.741 67.146 0.246 2.847
SSB: sum of squares between groups, SSW: sum of squares within groups, SST: sum of squares for total data,
F calculated: is the ratio of the mean square between groups to the mean square within groups, F critical: s the
critical value of the F-distribution for the chosen significance level (α= 0.05).
3.2.2. Chemical Characterisation
To verify the formation of thiourea linkage between FITC or RITC with APTES,
13
C
NMR was conducted on the reactants and the products. The focus of this analysis was on
the chemical shift of the carbon atom of the isothiocyanate group, which would convert to
a thiourea bond after reaction with the amino group of APTES, as illustrated in Figure 3.
Herein, we notice that the chemical shift value of the carbon atom of the isothiocyanate
group of FITC changed from (
δ
= 138.2,s) to (
δ
= 180.7,s) due to the formation of thiourea
with the amino group of APTES. Similar changes were observed for RITC, where the
value of the chemical shift of the carbon atom of the isothiocyanate group changed from
(
δ
= 137.9,s) to (179.3,s). NMR spectra for FITC and FITC-APTES, RITC and RITC-APTES
are demonstrated in Figures S4 and S5, respectively.

Biosensors 2024,14, 151 9 of 18
Biosensors2024,14,xFORPEERREVIEW9of19

Tab l e1.ANOVAparametersfortestingtheeffectofparticlesizevariabilityonthereproducibility
ofthefluorescencesignalforFMIPs,FNIPs,RMIPs,andRNIPs.
PolymerSSBSSWSSTFCalculatedFCritical(α=0.05)
FMIPs47.195180.125227.3210.6112.847
FNIPs3.61913.16316.7820.64152.847
RMIPs7.27224.67831.9500.6872.847
RNIPs6.40460.74167.1460.2462.847
SSB:sumofsquaresbetweengroups,SSW:sumofsquareswithingroups,SST:sumofsquaresfor
totaldata,Fcalculated:istheratioofthemeansquarebetweengroupstothemeansquarewithin
groups,Fcritical:sthecriticalvalueoftheF-distributionforthechosensignificancelevel(α=0.05).
3.2.2.ChemicalCharacterisation
ToverifytheformationofthiourealinkagebetweenFITCorRITCwithAPTES,13C
NMRwasconductedonthereactantsandtheproducts.Thefocusofthisanalysiswason
thechemicalshiftofthecarbonatomoftheisothiocyanategroup,whichwouldconvert
toathioureabondafterreactionwiththeaminogroupofAPTES,asillustratedinFigure
3.Herein,wenoticethatthechemicalshiftvalueofthecarbonatomoftheisothiocyanate
groupofFITCchangedfrom(δ=138.2,s)to(δ=180.7,s)duetotheformationofthiourea
withtheaminogroupofAPTES.SimilarchangeswereobservedforRITC,wherethevalue
ofthechemicalshiftofthecarbonatomoftheisothiocyanategroupchangedfrom(δ=
137.9,s)to(179.3,s).NMRspectraforFITCandFITC-APTES,RITCandRITC-APTESare
demonstratedinFiguresS4andS5,respectively.

Figure3.ThereactionbetweenFITCorRITCandAPTEStoformthefluorescentco-monomers.
Thesynthesisedpolymers,aswellastheorganicdyes,werecharacterisedviaIRspec-
troscopytoconfirmthecompleteinvolvementofthefluorescentdyesinthedeveloped
polymersandthespectraareshowninFigure4.BothFITCandRITCshowadistinctpeak
at~2010cm−1,whichischaracteristicoftheisothiocyanategroup.Asaresult,wenotice
thedisappearanceofthispeakintheresultingpolymers’spectra,whichverifiestheincor-
porationofthedyeintothepolymericstructure.Inaddition,weobservethatFMIPsand
FNIPs,aswellasRMIPsandRNIPs,havealmostidenticalspectrasincethereisnostruc-
turaldifferencebetweenthem.Therewerenopeaksintherangeof3000to3300cm−1,
Figure 3. The reaction between FITC or RITC and APTES to form the fluorescent co-monomers.
The synthesised polymers, as well as the organic dyes, were characterised via IR
spectroscopy to confirm the complete involvement of the fluorescent dyes in the developed
polymers and the spectra are shown in Figure 4. Both FITC and RITC show a distinct
peak at ~2010 cm
−1
, which is characteristic of the isothiocyanate group. As a result, we
notice the disappearance of this peak in the resulting polymers’ spectra, which verifies the
incorporation of the dye into the polymeric structure. In addition, we observe that FMIPs
and FNIPs, as well as RMIPs and RNIPs, have almost identical spectra since there is no
structural difference between them. There were no peaks in the range of 3000 to 3300 cm
−1
,
characteristic of amino and carboxylic groups, which indicates the complete removal of
pepsin from the binding sites of both MIPs.
Biosensors2024,14,xFORPEERREVIEW10of19

characteristicofaminoandcarboxylicgroups,whichindicatesthecompleteremovalof
pepsinfromthebindingsitesofbothMIPs.

Figure4.Infraredspectrafor(a)FITC,FMIPs,andFNIPsand(b)RITC,RMIPs,andRNIPs.
3.2.3.ThermalCharacterisation
DSCthermograms(Figure5(a1,a2))collectedfortheorganicdyesandtheirpolymers
providedfurtherconfirmationoftheincorporationofthedyesinthestructureofthede-
velopedpolymers.TheDSCcurveofFITCshowsonlyanexothermicpeakaround290°C,
yetDSCcurvesofbothFMIPsandFNIPsshowonlyonemeltingpeakataround320°C,
indicatingtheabsenceofanyresidualFITC.Similarly,theDSCthermogramforRITC
showedameltingpeakof210°C.However,theDSCgraphsofbothRMIPsandRNIPs
showedonlyonemeltingpeakat320°C,whichprovestheabsenceofanyresidualfluo-
rescentdye.ThisdatawasfurtherconfirmedwithTGA(Figure5(b1,b2)),whichshowed
nonotabledifferenceinthedecompositionpaernbetweenFMIPsandFNIPsandRMIPs
andRNIPsduetotheirstructuralsimilarity.However,theirdecompositionpaernsare
notablydifferentfromthatofthefluorescentdyes.
Figure 4. Infrared spectra for (a) FITC, FMIPs, and FNIPs and (b) RITC, RMIPs, and RNIPs.

Biosensors 2024,14, 151 10 of 18
3.2.3. Thermal Characterisation
DSC thermograms (Figure 5(a1,a2)) collected for the organic dyes and their polymers
provided further confirmation of the incorporation of the dyes in the structure of the
developed polymers. The DSC curve of FITC shows only an exothermic peak around
290
◦
C, yet DSC curves of both FMIPs and FNIPs show only one melting peak at around
320
◦
C, indicating the absence of any residual FITC. Similarly, the DSC thermogram for
RITC showed a melting peak of 210
◦
C. However, the DSC graphs of both RMIPs and
RNIPs showed only one melting peak at 320
◦
C, which proves the absence of any residual
fluorescent dye. This data was further confirmed with TGA (Figure 5(b1,b2)), which
showed no notable difference in the decomposition pattern between FMIPs and FNIPs and
RMIPs and RNIPs due to their structural similarity. However, their decomposition patterns
are notably different from that of the fluorescent dyes.
Biosensors2024,14,xFORPEERREVIEW11of19


Figure5.DSCthermogramsfor(a1)FITC,FMIPs,andFNIPs,(a2)RITC,RMIPs,andRNIPsand
TGAgraphsfor(b1)FITC,FMIPs,andFNIPs,(b2)RITC,RMIPs,andRNIPs.
3.2.4.FunctionalCharacterisation
TheresultsofthebindingkineticsexperimentsaregraphicallyshowninFigure6a,b.
Here,wenoticeadifferencebetweenfluorescein-dyedpolymersandrhodamineb-dyed
polymers.WeobservethatthepeakofbindingforFMIPstakesplaceafter2h;however,
forRMIPs,maximumbindingwasachievedafteronly1h.Wecanaributethefaster
bindingkineticsofRMIPstothepresenceofthetwodiethylaminogroupsinthestructure
ofrhodamineb,inwhichtheirpositivechargecaninteractsignificantlywiththenega-
tivelychargedpepsinmolecule,resultinginfasterbindingkinetics.Moreover,thevalue
ofQ(mgg
−1
)atthemaximumbindingtimeofRMIPs(256)ishigherthanthecorrespond-
ingQvalue(mgg
−1
)atthemaximumbindingtimeforFMIPs(217),whichagainverifies
theroleplayedbythediethylaminogroupsinenhancingthebindingofpepsin.Inan
aempttorecognisethemechanismofbindingofpepsintothedevelopedpolymers,
pseudo-first-orderandpseudo-second-orderkineticsparameterswereprocessedtofitthe
adsorptiondata.ThedatashowninTable2reflectsthatbothrhodaminebandfluorescein
imprintedpolymersfollowapseudo-second-ordermodel,whichconfirmsthatbinding
followsachemicaladsorptionmechanism.
Figure 5. DSC thermograms for (a1) FITC, FMIPs, and FNIPs, (a2) RITC, RMIPs, and RNIPs and TGA
graphs for (b1) FITC, FMIPs, and FNIPs, (b2) RITC, RMIPs, and RNIPs.
3.2.4. Functional Characterisation
The results of the binding kinetics experiments are graphically shown in Figure 6a,b.
Here, we notice a difference between fluorescein-dyed polymers and rhodamine b-dyed
polymers. We observe that the peak of binding for FMIPs takes place after 2 h; however,
for RMIPs, maximum binding was achieved after only 1 h. We can attribute the faster
binding kinetics of RMIPs to the presence of the two diethylamino groups in the structure
of rhodamine b, in which their positive charge can interact significantly with the negatively
charged pepsin molecule, resulting in faster binding kinetics. Moreover, the value of
Q (mg g
−1
) at the maximum binding time of RMIPs (256) is higher than the corresponding
Q value (mg g
−1
) at the maximum binding time for FMIPs (217), which again verifies the
role played by the diethylamino groups in enhancing the binding of pepsin. In an attempt
to recognise the mechanism of binding of pepsin to the developed polymers, pseudo-first-

Biosensors 2024,14, 151 11 of 18
order and pseudo-second-order kinetics parameters were processed to fit the adsorption
data. The data shown in Table 2reflects that both rhodamine b and fluorescein imprinted
polymers follow a pseudo-second-order model, which confirms that binding follows a
chemical adsorption mechanism.
Biosensors2024,14,xFORPEERREVIEW12of19


Figure6.Bindingkineticsfor(a)FMIPsandFNIPs,and(b)RMIPsandRNIPs,andbindingisotherm
for(c)FMIPsandFNIPs,and(d)RMIPsandRNIPs.
Tab l e 2.Bindingkineticsparametersfortheadsorptionofpepsinapplyingforpseudo-firstand
pseudo-secondbindingorders.
PseudoFirstOrderParameters
FMIPsFNIPs
K1(min−1)Qe(mgg−1)R2K1(min−1)Qe(mgg−1)R2
0.0007725.4320.00090.030981.1830.872
RMIPsRNIPs
K1(min−1)Qe(mgg−1)R2K1(min−1)Qe(mgg−1)R2
0.0338.110.660.017310.1750.163
PseudoSecondOrderParameters
FMIPsFNIPs
K2(gmg−1min−1)Qe(mgg−1)R2K2(gmg−1min−1)Qe(mgg−1)R2
0.0867196.090.99890.02809188.680.9961
RMIPsRNIPs
K2(gmg−1min−1)Qe(mgg−1)R2K2(gmg−1min−1)Qe(mgg−1)R2
0.0136156.250.98660.0221250.9759
K1andK2arethefirstandsecondorderrateconstantsrespectively,Qeisthequantityofpepsin
adsorbedpergramofpolymeratequilibrium,andR2isthelinearitycoefficient.
Thegraphicalrepresentationofthebindingisothermforbothfluoresceinandrhoda-
minebpolymers(Figure6c,d)showsalinearrelationshipbetweenthebindingcapacity
(Q)andtheconcentrationofpepsin.TheFreundlichisothermandtheLangmuirisotherm
modelswereappliedtothebindingisothermdata,andtheresultsareshowninTable3.
FromthecomputedR2values,wecanassumethatfluorescein-dyedpolymersfollowthe
Freundlichbindingisothermmodel,whichsuggeststhatthebindingisinmultiplelayers
onaheterogeneoussurface.Onthecontrary,rhodaminebdyedpolymersfollowedthe
Langmuirbindingmodel,whichindicatesthatbindingoccursinamonolayeroveraho-
mogenoussurface,whichagainprovesthesuperiorityofRMIPs.

Figure 6. Binding kinetics for (a) FMIPs and FNIPs, and (b) RMIPs and RNIPs, and binding isotherm
for (c) FMIPs and FNIPs, and (d) RMIPs and RNIPs.
Table 2. Binding kinetics parameters for the adsorption of pepsin applying for pseudo-first and
pseudo-second binding orders.
Pseudo First Order Parameters
FMIPs FNIPs
K1(min−1) Qe (mg g−1) R2K1(min−1) Qe (mg g−1) R2
0.00077 25.432 0.0009 0.0309 81.183 0.872
RMIPs RNIPs
K1(min−1) Qe (mg g−1) R2K1(min−1) Qe (mg g−1) R2
0.033 8.11 0.66 0.0173 10.175 0.163
Pseudo Second Order Parameters
FMIPs FNIPs
K2(g mg−1min−1) Qe (mg g−1) R2K2(g mg−1min−1) Qe (mg g−1) R2
0.0867 196.09 0.9989 0.02809 188.68 0.9961
RMIPs RNIPs
K2(g mg−1min−1) Qe (mg g−1) R2K2(g mg−1min−1) Qe (mg g−1) R2
0.0136 156.25 0.9866 0.022 125 0.9759
K
1
and K
2
are the first and second order rate constants respectively, Qe is the quantity of pepsin adsorbed per
gram of polymer at equilibrium, and R2is the linearity coefficient.
The graphical representation of the binding isotherm for both fluorescein and rho-
damine b polymers (Figure 6c,d) shows a linear relationship between the binding capacity
(Q) and the concentration of pepsin. The Freundlich isotherm and the Langmuir isotherm
models were applied to the binding isotherm data, and the results are shown in Table 3.

Biosensors 2024,14, 151 12 of 18
From the computed R
2
values, we can assume that fluorescein-dyed polymers follow the
Freundlich binding isotherm model, which suggests that the binding is in multiple lay-
ers on a heterogeneous surface. On the contrary, rhodamine b dyed polymers followed
the Langmuir binding model, which indicates that binding occurs in a monolayer over a
homogenous surface, which again proves the superiority of RMIPs.
Table 3. Adsorption isotherm parameters of MIPs and NIPs applying two models.
Langmuir Isotherm
FMIPs FNIPs
KL(L mg−1)
Q
max
(mg g
−1
)
RL R2KL(L mg−1)
Q
max
(mg g
−1
)
RL R2
0.002 19.45 1.002 0.9055 0.00158 25.22 1.001 0.9460
RMIPs RNIPs
KL(L mg−1)
Q
max
(mg g
−1
)
RL R2KL(L mg−1)
Q
max
(mg g
−1
)
RL R2
1.241 830.67 0.573 0.9887 1.226 174.093 3.782 0.9360
Freundlich Isotherm
FMIPs FNIPs
n KFR2n KFR2
0.381 349.869 0.9599 0.470 156.50 0.9428
RMIPs RNIPs
n KFR2n KFR2
1.285 553.384 0.9615 0.683 658.99 0.9117
K
L
and K
F
are the Langmuir constant and Freundlich constant, respectively. Q
max
is the theoretical maximum
adsorbed concentration, RL is the separation factor (1/1 + C
eq·
KL), n is the variation trend coefficient for the
adsorption isotherm, and R2is the linearity coefficient.
A binding selectivity assay towards pepsin was conducted for both FMIPs versus
FNIPs and for RMIPs versus RNIPs, and the imprinting factor (IF) was computed using
the formula.
IF =QMI Ps
QNIPs
Using a concentration of 1 mg mL
−1
of pepsin, the IF values were 1.13 for fluores-
cein dyed polymers and 1.89 for rhodamine b dyed polymers, which proved the higher
selectivity of RMIPs compared to FMIPs.
Furthermore, selectivity was tested against lipase and amylase due to their coexistence
with pepsin in human saliva along with lysozyme and thrombin as other proteins. The
results shown in Figure 7show remarkable selectivity towards pepsin in comparison to
other proteins for both fluorescein and rhodamine b dyed polymers, while again, RMIPs
showed superior selectivity, indicated by the values of the separation factor (
α
) calculated
using the formula.
α=QMI P·target
QMI P·competitor
3.3. Stability Testing
The results of the stability testing conducted over the period of 0-, 1-, 3-, and 6- months
showed a very slight decline in the binding capacity over the six month time period for both
FMIPs and RMIPs. Similar results were obtained for testing at low and high temperatures,
where the binding capacity for both FMIPs and RMIPs remained almost the same at all
temperatures. Results are depicted collectively in Figure 8. These results provide proof of
the superiority of MIPs to their corresponding antibodies in terms of physical stability.

Biosensors 2024,14, 151 13 of 18
Biosensors2024,14,xFORPEERREVIEW13of19

Tab l e 3.AdsorptionisothermparametersofMIPsandNIPsapplyingtwomodels.
LangmuirIsotherm
FMIPsFNIPs
KL(Lmg−1)Qmax(mgg−1)RLR2KL(Lmg−1)Qmax(mgg−1)RLR2
0.00219.451.0020.90550.0015825.221.0010.9460
RMIPsRNIPs
KL(Lmg−1)Qmax(mgg−1)RLR2KL(Lmg−1)Qmax(mgg−1)RLR2
1.241830.670.5730.98871.226174.0933.7820.9360
FreundlichIsotherm
FMIPsFNIPs
nKFR2nKFR2
0.381349.8690.95990.470156.500.9428
RMIPsRNIPs
nKFR2nKFR2
1.285553.3840.96150.683658.990.9117
KLandKFaretheLangmuirconstantandFreundlichconstant,respectively.Qmaxisthetheoretical
maximumadsorbedconcentration,RListheseparationfactor(1/1+Ceq·KL),nisthevariationtrend
coefficientfortheadsorptionisotherm,andR2isthelinearitycoefficient.
AbindingselectivityassaytowardspepsinwasconductedforbothFMIPsversus
FNIPsandforRMIPsversusRNIPs,andtheimprintingfactor(IF)wascomputedusing
theformula.
IF 𝑄
𝑄

Usingaconcentrationof1mgmL−1ofpepsin,theIFvalueswere1.13forfluorescein
dyedpolymersand1.89forrhodaminebdyedpolymers,whichprovedthehigherselec-
tivityofRMIPscomparedtoFMIPs.
Furthermore,selectivitywastestedagainstlipaseandamylaseduetotheircoexist-
encewithpepsininhumansalivaalongwithlysozymeandthrombinasotherproteins.
TheresultsshowninFigure7showremarkableselectivitytowardspepsinincomparison
tootherproteinsforbothfluoresceinandrhodaminebdyedpolymers,whileagain,
RMIPsshowedsuperiorselectivity,indicatedbythevaluesoftheseparationfactor(α)
calculatedusingtheformula.
𝛼 𝑄
𝑄 


Figure7.Selectivitystudiesof(a)FMIPsandFNIPsand(b)RMIPs,andRNIPsforpepsinagainst
otherproteins.

Figure 7. Selectivity studies of (a) FMIPs andFNIPs and (b) RMIPs, and RNIPs for pepsin against
other proteins.
Biosensors2024,14,xFORPEERREVIEW14of19

3.3.StabilityTesting
Theresultsofthestabilitytestingconductedovertheperiodof0-,1-,3-,and6-
monthsshowedaveryslightdeclineinthebindingcapacityoverthesixmonthtimepe-
riodforbothFMIPsandRMIPs.Similarresultswereobtainedfortestingatlowandhigh
temperatures,wherethebindingcapacityforbothFMIPsandRMIPsremainedalmostthe
sameatalltemperatures.ResultsaredepictedcollectivelyinFigure8.Theseresultspro-
videproofofthesuperiorityofMIPstotheircorrespondingantibodiesintermsofphysi-
calstability.

Figure8.StabilityofFMIPsandRMIPsagainsttimeandtemperature.
3.4.MechanismofFluorescenceQuenching
ToinvestigatethemechanismoffluorescencequenchingobservedforFMIPsand
RMIPsinthepresenceofpepsin,theStern-Volmerequationwasused.
𝐹0
𝐹1 𝐾𝑠𝑣󰇟𝑄󰇠
InwhichF0isthefluorescenceintensityofthefluorophorewithoutthequencher,F
isthefluorescenceintensityafterquenching,KsvistheStern-Volmerconstant,andQis
thequencherconcentration.Ideally,alinearrelationshipobservedintheStern-Volmer
plot(F0/F)versusquencherconcentration([Q])indicatesthedominanceofasingle
quenchingprocess(dynamicquenching).DeviationsfromlinearityinStern-Volmerplots
suggesttheinvolvementofdualquenchingmechanismssimultaneously,wheredynamic
quenchingandstaticquenchingplayroles[35,36].AsnoticedinFigure9,alinearrelation-
shipbetweenF0/Fandtheconcentrationofpepsin(thequencher)isobservedforboth
FMIPs(Figure9a)andRMIPs(Figure9b).Thislinearrelationshipreflectsadynamic
quenchingoffluorescenceinwhichthequencherinteractswiththefluorophoreatthe
excitedstateviacollisions,leadingtoareturntothegroundstatethroughnon-radiative
pathways.
Figure 8. Stability of FMIPs and RMIPs against time and temperature.
3.4. Mechanism of Fluorescence Quenching
To investigate the mechanism of fluorescence quenching observed for FMIPs and
RMIPs in the presence of pepsin, the Stern-Volmer equation was used.
F0
F=1+Ksv[Q]
In which F0 is the fluorescence intensity of the fluorophore without the quencher, Fis
the fluorescence intensity after quenching, Ksv is the Stern-Volmer constant, and Qis the
quencher concentration. Ideally, a linear relationship observed in the Stern-Volmer plot
(F0/F) versus quencher concentration ([Q]) indicates the dominance of a single quenching
process (dynamic quenching). Deviations from linearity in Stern-Volmer plots suggest the
involvement of dual quenching mechanisms simultaneously, where dynamic quenching
and static quenching play roles [
35
,
36
]. As noticed in Figure 9, a linear relationship between
F0/Fand the concentration of pepsin (the quencher) is observed for both FMIPs (Figure 9a)
and RMIPs (Figure 9b). This linear relationship reflects a dynamic quenching of fluorescence
in which the quencher interacts with the fluorophore at the excited state via collisions,
leading to a return to the ground state through non-radiative pathways.

Biosensors 2024,14, 151 14 of 18
Biosensors2024,14,xFORPEERREVIEW15of19


Figure9.GraphicalrepresentationofSternVolmerrelationshipbetweenconcentrationofpepsin
andthefluorescencequenchingfor(a)FMIPsand(b)RMIPs.
3.5.QuantitativeDetectionofPepsin
ThefluorescenceintensityofbothsolutionsofFMIPsandRMIPswasmeasuredin
triplicateattheirrespectiveexcitationwavelengths(λexc)afteraddingincreasedconcen-
trationsofpepsinfrom0(blank)to42.85µmolL−1.Itwasnotedthattheadditionofpepsin
causedaquenchinginfluorescenceintensityinaconcentration-dependentfashion,as
demonstratedinFigure10.However,athigherconcentrations,fluorescenceresponse
startedtoreachaplateau,whichindicatedthecompletesaturationofthebindingsitesfor
bothFMIPsandRMIPs.ThecalculatedlinearityparametersforbothMIPsareshownin
Tabl e 4.Interestingly,RMIPsshowedawiderlinearrangeextendingtoalowerconcen-
trationlimitof0.28µmolL−1andahigherlimitof42.85µmolL−1comparedtoFMIPs,
whoselowerlimitstartedat0.71µmolL−1,andthehigherlimitreached35.71µmolL−1.
Thisreinforcestheroleofthestructureofthefluorescentdyeinthebindingofthetarget
andthesensitivityoftheresultingMIPs.Therefore,RMIPshavemorecapabilitytobind
pepsinevenatlowerconcentrationscomparedtoFMIPsresultinginsuperiorsensitivity.
Inaddition,thelimitofdetectionvaluewascomputedforbothFMIPsandRMIPsusing
thefollowingequation:
LOD 3.3 𝛿
𝑠𝑙𝑜𝑝𝑒 
whereδisthestandarderroroftheinterceptoftheregressionequation.
ThecomputedLODvaluesfurtherconfirmedourhypothesis.WhereLODforFMIPs
is0.36µmolL−1,whilethevaluereached0.12µmolL−1forRMIPs.Inourstudy,wehave
achievedanLODof0.12µmolL−1forpepsinusingournovelfluorescentRMIPs.While
directcomparisonwithPeptest™intermsofconcentrationunitsmaynotbestraightfor-
wardduetodifferencesinassaymethodologies,ourMIP-basedapproachoffersacompa-
rablesensitivitytoconventionaldiagnostictestsforpepsindetection[25].Furthermore,
ourMIPsofferseveraladvantagesovertraditionalmethods,includingenhancedselectiv-
ity,easeofuse,andpotentialforintegrationintopoint-of-carediagnosticdevices.These
aributesmakeourMIPsapromisingtoolforsensitiveandspecificdetectionofpepsinin
variousbiologicalsamples,includingsaliva.
Figure 9. Graphical representation of Stern Volmer relationship between concentration of pepsin and
the fluorescence quenching for (a) FMIPs and (b) RMIPs.
3.5. Quantitative Detection of Pepsin
The fluorescence intensity of both solutions of FMIPs and RMIPs was measured in trip-
licate at their respective excitation wavelengths (
λexc
) after adding increased concentrations
of pepsin from 0 (blank) to 42.85
µ
mol L
−1
. It was noted that the addition of pepsin caused
a quenching in fluorescence intensity in a concentration-dependent fashion, as demon-
strated in Figure 10. However, at higher concentrations, fluorescence response started
to reach a plateau, which indicated the complete saturation of the binding sites for both
FMIPs and RMIPs. The calculated linearity parameters for both MIPs are shown in Table 4.
Interestingly, RMIPs showed a wider linear range extending to a lower concentration limit
of 0.28
µ
mol L
−1
and a higher limit of 42.85
µ
mol L
−1
compared to FMIPs, whose lower
limit started at 0.71
µ
mol L
−1,
and the higher limit reached 35.71
µ
mol L
−1
. This reinforces
the role of the structure of the fluorescent dye in the binding of the target and the sensitivity
of the resulting MIPs. Therefore, RMIPs have more capability to bind pepsin even at lower
concentrations compared to FMIPs resulting in superior sensitivity. In addition, the limit of
detection value was computed for both FMIPs and RMIPs using the following equation:
LOD =3.3·δ
slope
where δis the standard error of the intercept of the regression equation.
Table 4. Linearity parameters for quantitation of pepsin in standard solutions using FMIPs and RMIPs.
Parameter Linearity Range
(µmol L−1)Intercept ±SD aSlope ±SD bSyx cR2 d LOD (µmol L−1)e
FMIPs 0.71–35.71 551.94 ±1.61 −0.42 ±0.050 1.769 0.9842 0.36 ±0.051
RMIPs 0.28–42.85 307.53 ±0.85 −0.67 ±0.055 0.9733 0.9916 0.12 ±0.048
a
Standard deviation of the intercept,
b
Standard deviation of the slope,
c
Sum of square errors,
d
R the correlation
coefficient, eLOD limit of detection.
The computed LOD values further confirmed our hypothesis. Where LOD for FMIPs
is 0.36
µ
mol L
−1
, while the value reached 0.12
µ
mol L
−1
for RMIPs. In our study, we have
achieved an LOD of 0.12
µ
mol L
−1
for pepsin using our novel fluorescent RMIPs. While di-
rect comparison with Peptest™ in terms of concentration units may not be straightforward
due to differences in assay methodologies, our MIP-based approach offers a comparable
sensitivity to conventional diagnostic tests for pepsin detection [
25
]. Furthermore, our MIPs
offer several advantages over traditional methods, including enhanced selectivity, ease of

Biosensors 2024,14, 151 15 of 18
use, and potential for integration into point-of-care diagnostic devices. These attributes
make our MIPs a promising tool for sensitive and specific detection of pepsin in various
biological samples, including saliva.
Biosensors2024,14,xFORPEERREVIEW16of19
Figure10.(a)FluorescencemeasurementsforFMIPs,(b)calibrationcurveforFMIPsandFNIPs,(c)
fluorescencemeasurementsforRMIPs,and(d)calibrationcurveforRMIPsandRNIPs.
Tab l e 4.LinearityparametersforquantitationofpepsininstandardsolutionsusingFMIPsand
RMIPs.
ParameterLinearityRange(μmolL−1)Intercept±SDaSlope±SDbSyxcR2dLOD(μmolL−1)e
FMIPs0.71–35.71551.94±1.61−0.42±0.050 1.7690.98420.36±0.051
RMIPs0.28–42.85 307.53±0.85−0.67±0.055 0.97330.99160.12±0.048
aStandarddeviationoftheintercept,bStandarddeviationoftheslope,cSumofsquareerrors,dR
thecorrelationcoefficient,eLODlimitofdetection.
3.6.DevelopmentofaSeparation‐FreeQuantitationSystemforPepsininHumanSalivaSamples
ThefluorescentMIPswereeffectivelyappliedtoquantifypepsininhumansaliva
samples.Furthermore,duetothehighselectivityofthedevelopedFMIPsandRMIPs,
therewasnoneedforapriorextractionstep,thereforeeliminatinganextrastepinsample
preparationandallowingforasensitive,simple,andone-stepassay.Linearityinsaliva
rangedfrom1.42to42.85µmolL−1forRMIPsandfrom2.8to35.71µmolL−1forFMIPs.In
addition,thelowerconcentrationsmeasuredbyboththedevelopedMIPsareverysuita-
bleformeasuringtheverylowconcentrationsofpepsinfoundinthesalivaofrealGERD
patients.Moreover,thepercentageofrecoverywascalculatedforpepsinfromthreedif-
ferentspikedsamplescontainingthreedifferentconcentrationsalongthecalibration
range.Percentagesofrecoveryrangedfrom94.8to101.2%forFMIPsandfrom96.29to
100.21%forRMIPs;resultsaredemonstratedinTa b l e5.
Tab l e 5.Recoveryofspikedpepsinfromsalivasamples.
Concentration
(μmolmL−1)
AverageTotaltheAmountFound
(μmolmL−1)
Average%Recovery
±SDRSD%
FMIPsRMIPsFMIPsRMIPsFMIPsRMIPs
2.82.652.6994.84±3.3996.07±3.073.573.19
14.2814.2414.3199.76±1.51100.21±2.081.522.08
35.7136.1435.50101.22±0.31299.42±0.990.311.00
SDisthestandarddeviation,andRSDistherelativestandarddeviation.
Figure 10. (a) Fluorescence measurements for FMIPs, (b) calibration curve for FMIPs and FNIPs,
(c) fluorescence measurements for RMIPs, and (d) calibration curve for RMIPs and RNIPs.
3.6. Development of a Separation-Free Quantitation System for Pepsin in Human Saliva Samples
The fluorescent MIPs were effectively applied to quantify pepsin in human saliva
samples. Furthermore, due to the high selectivity of the developed FMIPs and RMIPs,
there was no need for a prior extraction step, therefore eliminating an extra step in sample
preparation and allowing for a sensitive, simple, and one-step assay. Linearity in saliva
ranged from 1.42 to 42.85
µ
mol L
−1
for RMIPs and from 2.8 to 35.71
µ
mol L
−1
for FMIPs.
In addition, the lower concentrations measured by both the developed MIPs are very
suitable for measuring the very low concentrations of pepsin found in the saliva of real
GERD patients. Moreover, the percentage of recovery was calculated for pepsin from three
different spiked samples containing three different concentrations along the calibration
range. Percentages of recovery ranged from 94.8 to 101.2% for FMIPs and from 96.29 to
100.21% for RMIPs; results are demonstrated in Table 5.
Table 5. Recovery of spiked pepsin from saliva samples.
Concentration
(µmol mL−1)
Average Total the Amount Found
(µmol mL−1)
Average% Recovery
±
SD
RSD %
FMIPs RMIPs FMIPs RMIPs FMIPs RMIPs
2.8 2.65 2.69 94.84 ±3.39 96.07 ±3.07 3.57 3.19
14.28 14.24 14.31 99.76 ±1.51
100.21
±
2.08
1.52 2.08
35.71 36.14 35.50 101.22 ±0.312 99.42 ±0.99 0.31 1.00
SD is the standard deviation, and RSD is the relative standard deviation.

Biosensors 2024,14, 151 16 of 18
4. Conclusions
In this work, we investigated the effect of the structure of the fluorescent dye on the
selectivity and sensitivity of fluorescent molecularly imprinted polymers. Two fluorescent
MIPs were prepared for the pepsin enzyme using two fluorescent dyes: fluorescein and rho-
damine b. Comparing the performance of both MIPs in terms of binding capacity, binding
selectivity, and quantitation range of pepsin confirmed the assumption that rhodamine b
can offer superior results owing to establishing two extra bonds with pepsin in comparison
to fluorescein. Thus, rhodamine b is assumed to provide three points of interaction with the
pepsin molecule. However, fluorescein provides only one point of interaction. This high-
lights the importance of applying a structure-based approach in the choice of fluorescent
dyes and monomers for the preparation of fluorescent MIPs. Furthermore, the developed
FMIPs and RMIPs were both applied successfully for the detection and quantitation of
pepsin in solutions and saliva samples, with RMIPs achieving higher sensitivity, indicated
by a LOD value of 0.12
µ
mol L
−1
compared to FMIPs with a LOD value of 0.36
µ
mol L
−1
.
Therefore, both fluorescent MIPs provided the advantage of an integrated extraction and
analysis tool in comparison to the magnetic MIPs our group previously developed, which
acted as extraction tools only.
Supplementary Materials: The following supporting information can be downloaded at: https://www.
mdpi.com/article/10.3390/bios14030151/s1, Figure S1. Pictures of the developed FMIPs, FNIPs, RMIPs,
and RNIPs on the bench and under long-wave UV light. Figure S2. Results of different optimisation
experiments for FMIPs and their corresponding FNIPs. Figure S3. Results of different optimisation
experiments for RMIPs and their corresponding RNIPs. Figure S4.
13
C NMR spectra of FITC and
FITC-APTES. Figure S5. 13C NMR spectra of RITC and RITC-APTES.
Author Contributions: Conceptualization, methodology, software, validation, formal analysis, inves-
tigation, resources, data curation, writing—original draft preparation, writing—review and editing,
and visualisation; A.M.M., supervision, project administration, funding acquisition; S.J.B., S.P.W. and
J.B. All authors have read and agreed to the published version of the manuscript.
Funding: This research was funded by Newton-Mosharafa Fund represented by The Egyptian Bureau
for Cultural and Educational Affairs in London and The APC was funded by Kingston University.
Institutional Review Board Statement: The study was conducted in accordance with the Declaration
of Helsinki, and approved by The Ethics Committee of Kingston University (code 2895, July 2022).
Informed Consent Statement: Informed consent was obtained from all subjects involved in the study.
Data Availability Statement: All data generated or analysed during this study are included in this
published article and its Supplementary Materials. Any additional data will be available upon request.
Acknowledgments: The authors kindly acknowledge the financial support offered by the Egyptian
Ministry of Higher Education and Scientific Research and The British Council (Newton-Mosharafa
Fund) represented by The Egyptian Bureau for Cultural and Educational Affairs in London. The
authors are also appreciative to Sigrid EyglóUnnarsdóttir and Boutaina Zbirta El Moujahid for their
precious help with this research.
Conflicts of Interest: The authors declare no conflicts of interest.
References
1.
Haupt, K.; Mosbach, K. Molecularly Imprinted Polymers and Their Use in Biomimetic Sensors. Chem. Rev. 2000,100, 2495–2504.
[CrossRef] [PubMed]
2.
Vasapollo, G.; Del Sole, R.; Mergola, L.; Lazzoi, M.R.; Scardino, A.; Scorrano, S.; Mele, G. Molecularly Imprinted Polymers:
Present and Future Prospective. Int. J. Mol. Sci. 2011,12, 5908–5945. [CrossRef] [PubMed]
3.
Quinn, T. Molecularly Imprinted Polymers (MIPS) Challenges, Uses and Prospects, 1st ed.; Nova Science Publishers, Inc.: Hauppauge,
NY, USA, 2016.
4.
Arabi, M.; Ostovan, A.; Li, J.; Wang, X.; Zhang, Z.; Choo, J.; Chen, L. Molecular Imprinting: Green Perspectives and Strategies.
Adv. Mater. 2021,33, 2100543. [CrossRef] [PubMed]
5.
Mostafa, A.M.; Barton, S.J.; Wren, S.P.; Barker, J. Review on Molecularly Imprinted Polymers with a Focus on Their Application to
the Analysis of Protein Biomarkers. TrAC Trends Anal. Chem. 2021,144, 116431. [CrossRef]

Biosensors 2024,14, 151 17 of 18
6.
Parisi, O.I.; Francomano, F.; Dattilo, M.; Patitucci, F.; Prete, S.; Amone, F.; Puoci, F. The Evolution of Molecular Recognition: From
Antibodies to Molecularly Imprinted Polymers (MIPs) as Artificial Counterpart. J. Funct. Biomater. 2022,13, 12. [CrossRef]
7.
Zhao, Y.; Chen, Y.; Fang, M.; Tian, Y.; Bai, G.; Zhuo, K. Silanized Carbon Dot-Based Thermo-Sensitive Molecularly Imprinted
Fluorescent Sensor for Bovine Hemoglobin Detection. Anal. Bioanal. Chem. 2020,412, 5811–5817. [CrossRef]
8. Fang, M.; Zhuo, K.; Chen, Y.; Zhao, Y.; Bai, G.; Wang, J. Fluorescent Probe Based on Carbon Dots/Silica/Molecularly Imprinted
Polymer for Lysozyme Detection and Cell Imaging. Anal. Bioanal. Chem. 2019,411, 5799–5807. [CrossRef]
9.
Piloto, A.M.L.; Ribeiro, D.S.M.; Rodrigues, S.S.M.; Santos, J.L.M.; Ferreira Sales, M.G. Label-Free Quantum Dot Conjugates for
Human Protein IL-2 Based on Molecularly Imprinted Polymers. Sens. Actuators B Chem. 2020,304, 127343. [CrossRef]
10.
Zhang, X.; Yang, S.; Sun, L.; Luo, A. Surface-Imprinted Polymer Coating l-Cysteine-Capped ZnS Quantum Dots for Target Protein
Specific Recognition. J. Mater. Sci. 2016,51, 6075–6085. [CrossRef]
11.
Zhou, T.; Halder, A.; Sun, Y. Fluorescent Nanosensor Based on Molecularly Imprinted Polymers Coated on Graphene Quantum
Dots for Fast Detection of Antibiotics. Biosensors 2018,8, 4944. [CrossRef]
12.
Piloto, A.M.; Ribeiro, D.S.M.; Rodrigues, S.S.M.; Santos, C.; Santos, J.L.M.; Sales, M.G.F. Plastic Antibodies Tailored on Quantum
Dots for an Optical Detection of Myoglobin down to the Femtomolar Range. Sci. Rep. 2018,8, 4944. [CrossRef] [PubMed]
13.
Pirzada, M.; Sehit, E.; Altintas, Z. Cancer Biomarker Detection in Human Serum Samples Using Nanoparticle Decorated
Epitope-Mediated Hybrid MIP. Biosens. Bioelectron. 2020,166, 112464. [CrossRef] [PubMed]
14.
Feng, H.; Wang, N.; Yuan, L.; Li, J.; Cai, Q. Surface Molecular Imprinting on Dye-(NH
2
)-SiO
2
NPs for Specific Recognition and
Direct Fluorescent Quantification of Perfluorooctane Sulfonate. Sens. Actuators B Chem. 2014,195, 266–273. [CrossRef]
15.
Wan, W.; Biyikal, M.; Wagner, R.; Sellergren, B.; Rurack, K. Fluorescent Sensory Microparticles That “Light-up” Consisting of a
Silica Core and a Molecularly Imprinted Polymer (MIP) Shell. Angew. Chem. Int. Ed. 2013,52, 7023–7027. [CrossRef] [PubMed]
16.
Xu, Z.; Deng, P.; Tang, S.; Li, J. Fluorescent Molecularly Imprinted Polymers Based on 1,8-Naphthalimide Derivatives for
Efficiently Recognition of Cholic Acid. Mater. Sci. Eng. C 2016,58, 558–567. [CrossRef] [PubMed]
17.
Sunayama, H.; Ohta, T.; Kuwahara, A.; Takeuchi, T. Fluorescence Signaling Molecularly Imprinted Polymers for Antibiotics
Prepared via Site-Directed Post-Imprinting Introduction of Plural Fluorescent Reporters within the Recognition Cavity. J. Mater.
Chem. B 2016,4, 7138–7145. [CrossRef] [PubMed]
18.
Wan, W.; Wagner, S.; Rurack, K. Fluorescent Monomers: “Bricks” That Make a Molecularly Imprinted Polymer “Bright”. Anal.
Bioanal. Chem. 2016,408, 1753–1771. [CrossRef]
19.
Chen, L.; Xu, S.; Li, J. Recent Advances in Molecular Imprinting Technology: Current Status, Challenges and Highlighted
Applications. Chem. Soc. Rev. 2011,40, 2922–2942. [CrossRef]
20. Liu, R.; Ko, C.C. Molecularly Imprinted Polymer-Based Luminescent Chemosensors. Biosensors 2023,13, 295. [CrossRef]
21.
Wang, F.; Wang, D.; Wang, T.; Jin, Y.; Ling, B.; Li, Q.; Li, J. A Simple Approach to Prepare Fluorescent Molecularly Imprinted
Nanoparticles. RSC Adv. 2021,11, 7732–7737. [CrossRef]
22.
Boulton, K.H.A.; Dettmar, P.W. A Narrative Review of the Prevalence of Gastroesophageal Reflux Disease (GERD). Ann. Esophagus
2022,5, 5–7. [CrossRef]
23. Samloff, I.M.; Taggart, R.T. Pepsinogens, Pepsins, and Peptic Ulcer. Clin. Investig. Med. 1987,10, 215–221.
24.
Wang, Y.J.; Lang, X.Q.; Wu, D.; He, Y.Q.; Lan, C.H.; Xiao, X.; Wang, B.; Zou, D.W.; Wu, J.M.; Zhao, Y.B.; et al. Salivary Pepsin as an
Intrinsic Marker for Diagnosis of Sub-Types of Gastroesophageal Reflux Disease and Gastroesophageal Reflux Disease-Related
Disorders. J. Neurogastroenterol. Motil. 2020,26, 74–84. [CrossRef] [PubMed]
25.
Zeleník, K.; Hránková, V.; Vrtková, A.; Staníková, L.; Komínek, P.; Formánek, M. Diagnostic Value of the Peptest
TM
in Detecting
Laryngopharyngeal Reflux. J. Clin. Med. 2021,10, 2996. [CrossRef] [PubMed]
26.
Verheyen, E.; Schillemans, J.P.; Van Wijk, M.; Demeniex, M.; Hennink, W.E.; Van Nostrum, C.F. Challenges for the Effective
Molecular Imprinting of Proteins. Biomaterials 2011,32, 3008–3020. [CrossRef] [PubMed]
27.
Ansari, S.; Masoum, S. Molecularly Imprinted Polymers for Capturing and Sensing Proteins: Current Progress and Future
Implications. TrAC Trends Anal. Chem. 2019,114, 29–47. [CrossRef]
28.
Pluhar, B.; Ziener, U.; Mizaikoff, B. Binding Performance of Pepsin Surface-Imprinted Polymer Particles in Protein Mixtures.
J. Mater. Chem. B 2015,3, 6248–6254. [CrossRef]
29.
Piletska, E.V.; Czulak, J.; Piletsky, S.S.; Guerreiro, A.; Canfarotta, F.; Piletsky, S.A. Novel Assay Format for Proteins Based on
Magnetic Molecularly Imprinted Polymer Nanoparticles—Detection of Pepsin. J. Chin. Adv. Mater. Soc. 2018,6, 341–351.
[CrossRef]
30.
García, Y.; Czulak, J.; Pereira, E.D.; Piletsky, S.A.; Piletska, E. A Magnetic Molecularly Imprinted Nanoparticle Assay (MINA) for
Detection of Pepsin. React. Funct. Polym. 2022,170, 105133. [CrossRef]
31.
Mostafa, A.M.; Barton, S.J.; Wren, S.P.; Barker, J. Development of Magnetic Molecularly Imprinted Polymers for the Extraction of
Salivary Pepsin Prior to Analysis by a Novel HPLC-SEC Method. Polymer 2022,261, 125417. [CrossRef]
32.
Wang, S.; Yin, D.; Wang, W.; Shen, X.; Zhu, J.J.; Chen, H.Y.; Liu, Z. Targeting and Imaging of Cancer Cells via Monosaccharide-
Imprinted Fluorescent Nanoparticles. Sci. Rep. 2016,6, 22757. [CrossRef]
33. Baliga, S.; Muglikar, S.; Kale, R. Salivary pH: A Diagnostic Biomarker. J. Indian Soc. Periodontol. 2013,17, 461–465. [CrossRef]
34. Fenger, F.; Andrew, R.H. On the Isoelectric Precipitation of Pepsin. J. Biol. Chem. 1927,73, 371–377. [CrossRef]

Biosensors 2024,14, 151 18 of 18
35.
Paterson, K.A.; Arlt, J.; Jones, A.C. Dynamic and Static Quenching of 2-Aminopurine Fluorescence by the Natural DNA
Nucleotides in Solution. Methods Appl. Fluoresc. 2020,8, 25002. [CrossRef]
36.
Gentleman, A.S.; Lawson, T.; Ellis, M.G.; Davis, M.; Turner-Dore, J.; Ryder, A.S.H.; Frosz, M.H.; Ciaccia, M.; Reisner, E.;
Cresswell, A.J.; et al. Stern-Volmer Analysis of Photocatalyst Fluorescence Quenching within Hollow-Core Photonic Crystal Fibre
Microreactors. Chem. Commun. 2022,58, 10548–10551. [CrossRef]
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual
author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to
people or property resulting from any ideas, methods, instructions or products referred to in the content.