|Year : 2020 | Volume
| Issue : 1 | Page : 35-40
Polymerase chain reaction for Clostridioides difficile infection detection: Necessity or redundancy? – A pilot study in a tertiary health-care centre in Central Kerala
Ansu Sam1, Seema Oommen2
1 Pushpagiri Institute of Medical Sciences and Research Centre, Thiruvalla, Kerala, India
2 Department of Microbiology, Lifecare Hospital, Musaffah, Abu Dhabi, UAE
|Date of Submission||02-Jan-2020|
|Date of Decision||24-May-2020|
|Date of Acceptance||26-Jun-2020|
|Date of Web Publication||13-Aug-2020|
Dr. Seema Oommen
Lifecare Hospital, Musaffah, Abu Dhabi
Source of Support: None, Conflict of Interest: None
INTRODUCTION: Clostridioides difficile infection (CDI) is one of the hospital-acquired infections and the most common cause for antibiotic-associated diarrhoea. Documentation of CDI is difficult, and interpretation of diagnostic results often requires consultation with clinical microbiologists. The purpose of this study was to compare the results of the combination of glutamate dehydrogenase enzyme (GDH) and toxin assay with the polymerase chain reaction (PCR) results, in order to find if the combination could substitute for the expensive molecular tests.
MATERIALS AND METHODS: The sample size was statistically calculated to be 30, using an SPSS software. Both GDH and toxin assay were simultaneously tested in all the randomly selected stool samples, by simple random sampling, and irrespective of the results, they were also tested for tcdB gene by PCR in the present study. All the samples were also plated onto Brazier's C. difficile agar and incubated anaerobically.
RESULTS: The sensitivity and specificity of GDH (using PCR as gold standard) were found to be 100% and 76.47%, respectively, and the sensitivity and specificity of toxin enzyme immunoassay (EIA) assay (using PCR as gold standard) were found to be 66.67% and 92.86%, respectively. However, when the toxin-equivocal results were also considered as positive, the sensitivity of toxin EIA was found to be 100%. The overall agreeability, using Cohen's Kappa statistic between GDH and toxin detection by enzyme-linked fluorescence assay, showed that they had moderate and substantial agreement, respectively, when compared to PCR.
CONCLUSIONS: In this study, each of the toxin negatives and positives was also PCR negative and positive, respectively. All the toxin-equivocal samples tested positive on PCR, so it is our conclusion that in the settings where they cannot be taken for further molecular testing, those samples be considered as harbouring toxigenic C. difficile.
Keywords: Clostridioides difficile infection, glutamate dehydrogenase, tcdB gene, toxin assay
|How to cite this article:|
Sam A, Oommen S. Polymerase chain reaction for Clostridioides difficile infection detection: Necessity or redundancy? – A pilot study in a tertiary health-care centre in Central Kerala. J Acad Clin Microbiol 2020;22:35-40
|How to cite this URL:|
Sam A, Oommen S. Polymerase chain reaction for Clostridioides difficile infection detection: Necessity or redundancy? – A pilot study in a tertiary health-care centre in Central Kerala. J Acad Clin Microbiol [serial online] 2020 [cited 2020 Oct 26];22:35-40. Available from: https://www.jacmjournal.org/text.asp?2020/22/1/35/292091
| Introduction|| |
There are few hurdles in medical diagnostics that remain equal across the globe and the difficulty in accurately diagnosing Clostridioides difficile infection (CDI) is one of them. The conundrum arises because each of the available tests looks for different targets, which makes it necessary to use an algorithmic or multistep approach unlike other infectious diseases where a single test is usually diagnostic.
CDI is caused by C. difficile, which until recently was known as Clostridium difficile. It is categorised as one of the hospital-acquired infections as it is the most common cause of antibiotic-associated diarrhoea and can easily get transmitted amongst hospitalised patients through the C. difficile spores that survive on fomites. Early diagnosis is of paramount importance to ensure that the disease is managed adequately.
However, documentation of C. difficile disease can be difficult. There are various commercially available immunoassays detecting C. difficile toxins that are simple and rapid, but recent studies have documented that these assays are insensitive (sensitivity <60%) and in some cases non-specific., The tests can detect one or both of the toxins and/or the glutamate dehydrogenase (GDH) enzyme. As some C. difficile strains do not produce toxin A, it is recommended that toxin enzyme immunoassays (EIAs) should detect both toxins A and B., GDH is a cytoplasmic enzyme and the GDH detection assays detect both toxigenic and non-toxigenic C. difficile strains. For this reason, GDH is only recommended to be used in combination with other assays, such as a toxin immunoassay.
Nucleic acid amplification test (NAAT) detecting the tcdB gene is the definitive diagnostic modality, with a turnaround time of 1–2 h and higher sensitivity and specificity (93.2% and 96.9% respectively) than the EIAs, but it requires expertise and is not currently economical for most patients. Studies have also shown that exclusively relying on molecular tests leads to overdiagnosis since polymerase chain reaction (PCR) looks for the gene responsible for toxin production and not the toxins itself. Therefore, the case scenario where colonisation of toxigenic C. difficile has occurred without toxin production can be falsely diagnosed as CDI if toxin EIA is not done. An alternative approach used in some laboratories is to screen stool specimens for GDH and then confirm the presence of toxin-producing C. difficile using the cytotoxicity assay or NAATs for the toxins.
The Society for Healthcare Epidemiology of America-Infectious Diseases Society of America (SHEA-IDSA) guidelines, 2018, recommended a multistep algorithm using GDH as a screening test plus toxin or GDH plus toxin arbitrated by a PCR or nucleic acid detection plus toxin rather than a nucleic acid testing alone. GDH is recommended as the initial screening step in the two- or three-step algorithm as it is a very sensitive test detecting the highly conserved metabolic enzyme that is present in high levels in all isolates of C. difficile, both toxigenic and non-toxigenic strains. Combining the GDH test with toxin assay makes the test more specific and yields rapid results. As recommended by the SHEA-IDSA, this combination can then be arbitrated by PCR testing as needed, which makes the algorithm much more economical. In a resource-limited setting as ours that cater to patients from lower socioeconomic status, an expensive test like PCR is not financially feasible as a routine option. The purpose of this pilot study is to test a small but statistically valid number of samples to compare the diagnostic results of combining GDH and toxin assay with those of PCR.
| Materials and Methods|| |
This was a cross-sectional study conducted in the Department of Microbiology, from October 2016 to October 2018, in a 1200-bedded tertiary care hospital in Central Kerala, India, after obtaining institutional ethical clearance (PIMSRC/E1/388A/50/2016). Assuming a sensitivity of 95% and specificity of 95% for the combination of EIA for C. difficile toxin with GDH detection tests along with an expected prevalence of 18% and an alpha error of 5%, the minimum sample size was calculated to be 30, with an SPSS software using the formula n = 4pq/d 2. Consecutive sampling method was employed and informed consent was taken. Faecal samples of patients with sudden-onset diarrhoea (that was not present at that time of admission and which followed a history of antibiotic therapy) were included in the study. Previous antimicrobial drug therapy was defined as the receipt of an oral or parenteral antimicrobial agent for >72 h within the preceding two months. All formed stool samples as well as those who had diarrhoea due to other known causes such as infective gastroenteritis and taking laxatives were excluded from the study.
All the samples were screened for the presence of C. difficile GDH enzyme and toxin A and B by enzyme-linked fluorescence assay (ELFA) and VIDAS (BioMerieux). The positive cut-off value for C. difficile GDH assay was >0.10 IU/mL and for C. difficile toxin A and B assay was >0.37 IU/mL. The measures taken by the bacteriology laboratory including reporting were subjective to the results obtained from this dual testing. Those samples that were positive in both tests were reported as toxigenic C. difficile to the treating physician and the infection control nurse. Samples that were positive for GDH enzyme but negative for toxins were reported as non-toxigenic C. difficile and PCR was suggested as an additional test to confirm diagnosis with an alert sent to the infection control team. This was done because even a non-toxigenic C. difficile shedder could release spores that could affect the other vulnerable patients who were in close proximity. Proper measures such as isolation or cohorting were done as deemed fit by the infection control team. Those that were negative for GDH enzyme and C. difficile toxin were reported as negative for CDI.
Both GDH and toxin assay were simultaneously tested in all the randomly selected samples, and irrespective of the results, they were also tested for tcdB gene by PCR in the present study. Toxin A and toxin B were coded for by tcdA and tcdB, respectively. While toxin A was enterotoxic, toxin B was cytotoxic, and studies have shown that though in most cases, both the toxins were present, there were many cases where toxin A is absent but toxin B has been detected in all cases diagnosed as CDI. Studies have shown that the gene tcdB-producing toxin B was found to be more significant in disease pathogenesis. Therefore, in this study, the target for molecular testing was selected as tcdB gene.
All the samples were also plated onto Brazier's C. difficile agar with 1% lysed sheep blood and incubated anaerobically for seven days and intermittently checked for growth on fifth and seventh day. Growth supplement and egg yolk emulsion were also added to the media to inhibit the growth of other bacteria as well as to differentiate C. difficile from other lecithinase-positive Clostridia. The anaerobic environment for the medium was provided in an anaerobic jar along with a GasPak pouch. Growth was examined for characteristic appearance, smell and Gram stain morphology. Grey, opaque, flat-raised, circular to elongate 4–6 mm colonies with horse stable odour and showing Gram positive to variable bacilli with subterminal spores were presumptively identified as C. difficile. The stool samples were cultured only after subjecting them to alcohol shock, by aliquoting 0.5 ml of stool, vortexed briefly with 1 ml of 70% ethanol, incubated for one hour at room temperature to eliminate all the non-sporing organisms found in stool. The anaerobic environment ensured that all the aerobic and facultative anaerobic organisms did not survive the culture. The confirmatory tests for C. difficile isolation is Cytotoxigenic Culture or Cytotoxicity Assay, however both the tests were outside the scope of this study and were not done. C. difficile was identified by eliminating the possibility of other organisms by using selective growth media. The lecithinase-negative property of the colonies eliminated the possibility of C. perfringens which can cause diarrhoea by food poisoning.
| Results|| |
Out of the 30 test samples, 17 were GDH positive and the remaining 13 were GDH negative [Figure 1]. Amongst the GDH positives, nine were toxin positive and four were toxin equivocal and the remaining were negative. All nine toxin positive and the four toxin-equivocal tested positive on PCR analysis. Only 11 samples out of the 30 yielded growth, which were greyish white opaque, flat, rough-edged colonies with a wax-like consistency with no lecithinase activity [Figure 2].
|Figure 1: Results for glutamate dehydrogenase and toxin enzyme immunoassay|
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| Discussion|| |
Amongst the selected study samples (n = 30), there were nine that were GDH and toxin positive, while eight were positive for GDH but tested negative or equivocal for toxin production. Four out of these eight GDH-positive samples had an equivocal toxin result [Figure 1]. The remaining 13 samples were GDH as well as toxin negative.
Thus, 17 samples were positive for GDH, of which the nine that were toxin positive by ELFA had GDH values ranging from 3.96 to 8.33 IU, with a mean of 6.75 IU [Figure 3]. The GDH values amongst the GDH positives with toxin negative (n = 4) ranged from 1.5 IU to 11.7 IU.
|Figure 3: Values of glutamate dehydrogenase amongst toxin positives (n = 9), negatives (n = 4) and equivocal results (n = 4); where X-axis shows the study sample and Y-axis shows the glutamate dehydrogenase value in IU/ml|
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The patient with the highest GDH value of 11.7 had negative toxin result. This was a 67-year-old male who is a known case of chronic kidney disease with a positive sputum culture with K. pneumoniae and treated with Meropenem two weeks before testing for CDI detection. Although this patient was non-toxigenic on testing, he was thought to be a carrier, who could potentially affect other patients. IDSA recommends that such cases be considered as “excretors”, who do not require treatment but present an infection control risk. Retesting in CDI detection is not recommended because multiple repeat testing within seven days runs a high risk of generating false-positive results. Repeat testing must be done only if diarrhoea persists and/or if symptoms worsen. In a previous study done in our hospital, we found that isolating with barrier precautions even those patients who are non-toxigenic with positive GDH significantly reduced the prevalence of CDI in one year. The case was thought to be toxin negative due to toxin inhibition in the patient or due to the below detectable levels of toxin as suggested by studies like Riggs et al. Since the rest of the toxin-negative GDH positives had GDH values that were at or below 7 IU, it was assumed that the toxin assay may have been false negative and PCR would turn out to be positive in this case. However, this was deemed a significant case in our study when the PCR was also yielded a negative result as it concurred with the non-toxigenic result, making the combination of GDH and toxin assay a valid one.
The GDH values of the four equivocal toxin samples were 10.3, 5.12, 3.12 and 1.66 IU. The equivocal strains were evidently found to be toxin producers by subjecting them to toxin B gene detection by PCR, thus bringing the total toxin producing strains amongst the random samples (n = 30) from nine to 13 with a GDH mean value of 6.38 IU amongst them. Previous studies have shown that most of the samples with an equivocal toxin result were positive by PCR.,, A study published in the UK in 2017 by Kumar et al. found that in a 207-sample study, 92 samples were detected to be toxin positive by PCR but negative EIA. However, they found that a positive toxin EIA is a significant independent predictor of death with an odd ratio of 1.89, while PCR positivity in negative EIA is not, which means that, because the sensitivity of PCR is very high (100% with respect to cytotoxicity assays), as shown by a meta analysis study, more cases will be detected by PCR, which had been negative by other assays. However, regardless of the PCR result, if toxin was negative by EIA, the patient outcome may not be affected. Amongst the four GDH-positive and toxin ELFA-negative samples, the GDH mean was found to be 5.92 IU.
There are several studies which consider the GDH positivity with their values for toxin-negative samples.,, A study done by Davies et al., 2015, found that out of the 1428 GDH-equivocal samples by ELFA, 424 were positive by PCR. In the present study, we have optical density values for GDH-equivocal results since ELFA was used, but in most of the studies, the GDH test is a lateral flow assay. Regardless of the method of detection, the main suggestion across literature is to test these discrepant samples for PCR testing., However, in resource-limited setting as ours, we can use the values from the present study as pointers whether a further PCR evaluation is necessary. It is a suggestion from this study that since the lowest GDH positive value for toxin positives was 3.96 IU, PCR need not be done for those toxin-negative samples with a GDH value of lower than 3 IU. CDI prevalence studies have also shown this to be true where toxin-negative samples that are GDH positive usually have a lower value compared to the toxin-positive samples. However, regardless of the GDH value, if a toxin result is equivocal, the sample must be tested by molecular methods or be considered as toxigenic if PCR cannot be done. The National Health Services of Scotland guidelines says that in view of a discrepant result, like GDH result positive and Toxin result below detection level by immunoassay the test must be repeated, and clinical correlation must also be made. If the patient is symptomatic, the laboratory result must not be the only reason why treatment should be withheld.
The sensitivity and specificity of GDH (using PCR as gold standard) were found to be 100% and 76.47%, respectively, with a confidence interval of 95% and the sensitivity and specificity of toxin EIA assay (using PCR as gold standard) were found to be 66.67% and 92.86%, respectively, with a confidence interval of 95%. However, when the toxin-equivocal results were also considered as positive, the sensitivity of toxin EIA was found to be 100% [Table 1]. The overall agreeability between GDH and toxin detection by ELFA showed that they had moderate and substantial agreement, respectively, when compared to PCR [Table 2].,, This shows that the combination of GDH and toxin assay can adequately substitute for molecular detection of tcdB gene by PCR.
|Table 1: Comparison of glutamate dehydrogenase enzyme assay and toxin assay with respect to polymerase chain reaction test (toxin 9 positives and 4 equivocal)|
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|Table 2: Agreement of glutamate dehydrogenase enzyme assay and toxin assay results between polymerase chain reaction test|
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All the samples were also subjected to anaerobic incubation, where, true to its name, C. difficile proved to be difficult to culture. Only 11 samples out of the 30 yielded growth, which were greyish white opaque, flat, rough-edged colonies with a wax-like consistency with no lecithinase activity [Figure 2]. It also produced a characteristic horse barn smell of C. difficile when the plates were opened. Gram stain showed Gram variable bacilli with bulging spores  [Figure 4]. However, the culture took as long as six–seven days to yield growth and thus CDI cannot be diagnosed solely by culture and morphology.,
|Figure 4: Gram staining of smear from the colony Gram variable bacilli with bulging spores. Image on the right shows magnified view of spores|
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One of the limitations of the study is that it is single centric with a relatively small sample size. A larger number of study samples would have been ideal, but the high cost of the PCR inclusive of the DNA extraction for each sample was the compelling reason for scaling down. However, this size was statistically sound and taken as per the calculated minimum required sample size to make this study valid.
| Conclusions|| |
Out of the three clinically significant tests for CDI detection, namely PCR, GDH assay and toxin assay, the most reliable as well as economical testing method is the combination of GDH and toxin assay [Table 3]. In this study, each of the toxin negatives and positives were also PCR negative and positive, respectively. All the toxin-equivocal samples tested positive on PCR, so it is our conclusion that in the settings where they cannot be taken for further molecular testing, those samples be considered as harbouring toxigenic C. difficile. Culture yielded characteristic growth after prolonged incubation, with a minimum of five days, which negated the purpose of employing it as a clinical diagnostic method.
|Table 3: Comparison pros and cons between glutamate dehydrogenase enzyme, toxin enzyme immunoassays and polymerase chain reaction|
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Conflicts of interest
There are no conflicts of interest.
| References|| |
Mullish BH, Williams HR. Clostridium difficile
infection and antibiotic-associated diarrhoea. Clin Med (Lond) 2018;18:237-41.
McDonald LC, Gerding DN, Johnson S, Bakken JS, Carroll KC, Coffin SE, et al
. Clinical practice guidelines for Clostridium difficile
infection in adults and children: 2017 update by the Infectious Diseases Society of America (IDSA) and Society for Healthcare Epidemiology of America (SHEA). Clin Infect Dis 2018;66:e1-48.
Burnham CA, Carroll KC. Diagnosis of Clostridium difficile
infection: An ongoing conundrum for clinicians and for clinical laboratories. Clin Microbiol Rev 2013;26:604-30.
Tenover FC, Baron EJ, Peterson LR, Persing DH. Laboratory diagnosis of Clostridium difficile
infection can molecular amplification methods move us out of uncertainty? J Mol Diagn 2011;13:573-82.
Fang FC, Polage CR, Wilcox MH. Point-counterpoint: What is the optimal approach for detection of Clostridium difficile
Infection? J Clin Microbiol 2017;55:670-80.
Goldenberg SD, Cliff PR, Smith S, Milner M, French GL. Two-step glutamate dehydrogenase antigen real-time polymerase chain reaction assay for detection of toxigenic Clostridium difficile
. J Hosp Infect 2010;74:48-54.
Martínez-Meléndez A, Camacho-Ortiz A, Morfin-Otero R, Maldonado-Garza HJ, Villarreal-Treviño L, Garza-González E. Current knowledge on the laboratory diagnosis of Clostridium difficile
infection. World J Gastroenterol 2017;23:1552-67.
Humphries RM, Uslan DZ, Rubin Z. Performance of Clostridium difficile
toxin enzyme immunoassay and nucleic acid amplification tests stratified by patient disease severity. J Clin Microbiol 2013;51:869-73.
Polage CR, Gyorke CE, Kennedy MA, Leslie JL, Chin DL, Wang S, et al
. Overdiagnosis of Clostridium difficile
infection in the molecular test era. JAMA Intern Med 2015;175:1792-801.
Crobach MJ, Planche T, Eckert C, Barbut F, Terveer EM, Dekkers OM, et al
. European Society of clinical microbiology and infectious diseases: Update of the diagnostic guidance document for Clostridium difficile
infection. Clin Microbiol Infect 2016;22 Suppl 4:S63-81.
Lee NY, Huang YT, Hsueh PR, Ko WC. Clostridium difficile
bacteremia, taiwan1. Emerg infect Dis 2010;16:1204.
Claro T, Daniels S, Humphreys H. Detecting Clostridium difficile
spores from inanimate surfaces of the hospital environment: Which method is best? J Clin Microbiol 2014;52:3426-8.
Carter GP, Awad MM, Kelly ML, Rood JI, Lyras D. TcdB or not TcdB: A tale of two Clostridium difficile
toxins. Future Microbiol 2011;6:121-3.
Sam AS, Jacob AM, Oommen S. Prevalence and outcome of clostridioides difficile infection in a tertiary care hospital in Kerala, India. National Journal of Laboratory Medicine 2019;8:MO01-4.
Riggs MM, Sethi AK, Zabarsky TF, Eckstein EC, Jump RL, Donskey CJ. Asymptomatic carriers are a potential source for transmission of epidemic and nonepidemic Clostridium difficile
strains among long-term care facility residents. Clin Infect Dis 2007;45:992-8.
Eastwood K, Else P, Charlett A, Wilcox M. Comparison of nine commercially available Clostridium difficile
toxin detection assays, a real-time PCR assay for C. difficile
tcdB, and a glutamate dehydrogenase detection assay to cytotoxin testing and cytotoxigenic culture methods. J Clin Microbiol 2009;47:3211-7.
Brown NA, Lebar WD, Young CL, Hankerd RE, Newton DW. Diagnosis of Clostridium difficile
infection: Comparison of four methods on specimens collected in Cary-Blair transport medium and tcdB PCR on fresh versus frozen samples. Infect Dis Rep 2011;3:e5.
Davies KA, Berry CE, Morris KA, Smith R, Young S, Davis TE, et al
. Comparison of the Vidas C. difficile
GDH automated enzyme-linked fluorescence immunoassay (ELFA) with another commercial enzyme immunoassay (EIA) (Quik Chek-60), Two selective media, and a PCR assay for gluD for detection of Clostridium difficile
in fecal samples. J Clin Microbiol 2015;53:1931-4.
Cheng JW, Xiao M, Kudinha T, Xu ZP, Sun LY, Hou X, et al
. The role of glutamate dehydrogenase (GDH) testing assay in the diagnosis of Clostridium difficile
infections: A high sensitive screening test and an essential step in the proposed laboratory diagnosis workflow for developing countries like China. PLoS One 2015;10:e0144604.
Planche T, Wilcox MH. Diagnostic pitfalls in Clostridium difficile
infection. Infect Dis Clin North Am 2015;29:63-82.
Davies KA, Ashwin H, Longshaw CM, Burns DA, Davis GL, Wilcox MH, EUCLID Study Group. Diversity of Clostridium difficile PCR ribotypes in Europe: results from the European, multicentre, prospective, biannual, point-prevalence study of Clostridium difficile infection in hospitalised patients with diarrhoea (EUCLID), 2012 and 2013. Eurosurveillance. 2016;21:30294.
Murad YM, Perez J, Ybazeta G, Mavin S, Lefebvre S, Weese JS, et al
. False negative results in Clostridium difficile
testing. BMC Infect Dis 2016;16:430.
Qutub M, Govindan P, Vattappillil A. Effectiveness of a Two-Step Testing Algorithm for Reliable and Cost-Effective Detection of Clostridium difficile
Infection in a Tertiary Care Hospital in Saudi Arabia. Med Sci (Basel). 2019;7:6. Published 2019 Jan 8. doi:10.3390/medsci7010006.
Scottish Health Protection Network. Guidance on Prevention and Control of Clostridium difficile Infection (CDI) in health and social care settings in Scotland. Health Protection Network Scottish Guidance. (2017 Edition). Health Protection Scotland, Glasgow, 2017.
Curry SR. Clostridium difficile
. Clin Lab Med 2017;37:341-69.
Peterson LR, Mehta MS, Patel PA, Hacek DM, Harazin M, Nagwekar PP, et al
. Laboratory testing for Clostridium difficile
infection: Light at the end of the tunnel. Am J Clin Pathol 2011;136:372-80.
Yoldaş Ö, Altındiş M, Cufalı D, Aşık G, Keşli R. A diagnostic algorithm for the detection of Clostridium difficile
-associated diarrhea. Balkan Med J 2016;33:80-6.
Deshpande A, Pasupuleti V, Rolston DD, Jain A, Deshpande N, Pant C, et al
. Diagnostic accuracy of real-time polymerase chain reaction in detection of Clostridium difficile
in the stool samples of patients with suspected Clostridium difficile
Infection: A meta-analysis. Clin Infect Dis 2011;53:e81-90.
[Figure 1], [Figure 2], [Figure 3], [Figure 4]
[Table 1], [Table 2], [Table 3]